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Deciphering the role of antherozoid specific DNA
methyltransferases in Physcomitrella patens
Sónia Alexandra Gomes Pereira
Thesis to obtain the Master of Science Degree in
Biotechnology
Supervisors: Doctor Leonilde de Fátima Morais Moreira
and Doctor Jörg-Dieter Becker
Examination Committee
Chairperson: Doctor Miguel Nobre Parreira Cacho Teixeira
Supervisor: Doctor Jörg-Dieter Becker
Members of the Committee: Doctor Maria Wanda Sarujine Viegas
November, 2015
“All we have to decide is what to do with the time that is given us.”
- Gandalf,
J.R.R. Tolkien,
Fellowship of the Ring, Second chapter,
"The Shadow of the Past",
For Liz,
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Acknowledgements
I acknowledge Instituto Gulbenkian de Ciência for the opportunity to conduct this work. I also want
to show my gratitude to my supervisors Doctor Jörg Becker and Doctor Leonilde Moreira for the
professional and personal support during the progression of this work.
I am grateful to all my co-workers for all the knowledge and support during my time in the
laboratory, particularly to Marcela Coronado for teaching me the basis about working with the wonderful
and tiny moss Physcomitrella patens and to Leonor Boavida for always being available to help me and
the indispensable help with the cloning (even on weekends or late hours). I also thank Mário Santos for
the help in obtaining some colony area images, Anna Thamm for the generation of the pAT05 plasmid
and the Δdrm2 lines, Ann-Cathrin Lindner for all her lovely energy and advices in these past few months,
Patricia Pereira for all the company and wonderful conversations in the lab as well as Joana Caria and
Custódio Nunes for all the good times in the lab.
I would also like to acknowledge all the help of the technicians of the Genomics and Gene
Expression units of the IGC: Susana Ladeiro, João Costa and João Sobral for their help in the
sequencing of the pAT05 plasmid and the sequencing of the flanking regions of the cloning and also to
Daniel Sobral of the Bioinformatics unit for the help with the NGS data assembly. I would also like to
point out Nuno Moreno’s (IGC’s imaging facility) help in developing the process to treat the colony area
data and Cláudia Bispo’s availability, enthusiasm and help in the flow cytometric analysis of the
antherozoid samples!
I appreciate all the advices related to phylogeny given to me Stefan Rensing, Magdanela
Bezanilla’s advices about the transformation protocol and all my friends at the IGC for making it such a
wonderful place to work!
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Agradecimentos
Este ano foi uma autêntica montanha russa, cheia de altos, baixos e loops. Foram várias as
vezes que duvidei chegar aqui, ao fim deste ciclo.... Agora é meu dever agradecer a todos os que me
impediram de desistir e me possibilitaram chegar aqui.
Primeiramente quero agradecer todo o apoio do meu grupo de laboratório Plant Genomics e do
meu orientador Jörg Becker, particularmente à Leonor pela ajuda essencial nas clonagens e pela
companhia de sábado à tarde.... Quero também agradecer aos meus compinchas de almoço: João
Sobral, João Costa e Susana Ladeiro por todas as horas passadas a reclamar, desabafar, brincar,
debater e ter conversas parvas, mas também a trabalhar claro!
Agradeço à minha família todo o apoio ao longo deste longo ano e toda a paciência e
preocupação que demonstraram, especialmente aos meus avós maternos, irmã, mãe e Jacinto que
sempre lá estiveram quando mais precisei.
À minha gata – Liz, à qual dedico este trabalho e que, infelizmente não aguentou para me ver
terminá-lo, mas que sempre esteve sempre ao meu lado (e ao meu colo, e na minha barriga... etc..) e
cuja presença me fazia esquecer o mundo... A sua falta será sempre sentida.
Obviamente um grande, enorme e gigante Obrigado ao meu namorado, Miguel, por sempre ter
estado a meu lado apesar de todas as dificuldades que surgiram ao longo deste ano. Não exagero ao
dizer que sem ti tenho a certeza que não estaria aqui!
Às minhas incríveis amigas e compinchas de sugar-crushes: Adriana, Joana e Leonor por todos
os momentos em que o açúcar melhorou as nossas vidas nos últimos largos meses e por todas as
lamentações que ouviram. E claro, pelo vosso carinho e preocupação. Aos meus guerreiros Hwarang,
em particular à Catarina, Eiras e o fantástico Sambonim o meu mais sincero obrigado por me motivarem
a regressar a casa, puxarem por mim e me ajudarem a manter a pouca sanidade mental que ainda me
sobra! Mas também aos restantes membros do grupo que me motivam para continuar e nunca desistir.
Infelizmente sou também obrigada a agradecer a toda a equipa de médicos, enfermeiros e
assistentes de saúde que me acompanharam nalguma fase deste longo processo. Em especial ao Dr.
Miguel Rebocho e ao Dr. Manuel Cunha e Sá por me salvarem a vida.
Obrigada também aos Oliveira, Lourenço, Bento, Martins.... Obrigada a todos os meus
professores, orientadores, colegas que sempre me estimularam a querer saber, procurar e responder
e que me levaram a este momento. E obrigada pelo especial carinho que recebo de grande parte deles
quando os vejo.
A todos os que estiverem de algum modo presentes na minha vida ao longo desta fase, que me
felicitaram nos bons momentos, que me ajudaram nos piores e que me deram nas orelhas nos
momentos de angústia... A todos os que partilharam os bons e os maus momentos, cafés, cervejas,
açúcar, saladas (felizmente não muitas!), pontapés, suor, sangue, lágrimas e sorrisos.
O meu mais sincero obrigado a todos vós! Espero que todos possam continuar a estar presentes
na próxima fase da minha vida porque a minha vida não seria a mesma sem vós!
Obrigada,
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Abstract
Cytosine methylation represents the most common DNA modification in eukaryotic genomes. In
plants 5-mC can be detected in CG, CHG and CHH contexts, being catalyzed by DNA
methyltransferases. In antherozoids of Physcomitrella patens expression of de novo DNA
methyltransferases is limited to PpDRM2 and PpDNMT3b, standing in stark contrast with the broad
expression of other DNA methyltransferases during the life cycle of this model bryophyte. Given the
importance of DNA methylation for genome integrity this observation prompted us to study the role of
de novo methylation during sexual reproduction in Physcomitrella.
We obtained two independent Δdrm2 knockout lines and analyzed their fertilization rates in
comparison to the wild-type. In the F0 lower rates were detected for Δdrm2#1, but not for Δdrm2#2. F1
and F2 lines showed no variation in rates, indicating a possible compensatory mechanism after the first
fertilization. Based on an observation that prolonged cold storage of spores led to irregular shaped
colonies with gametophores in wild-type and smaller and round colonies lacking gametophores in
Δdrm2, we performed a time-course phenotyping experiment. To this end methods for high-throughput
colony area and dry weight assessment were established. The variation in colony growth was observed
again, but it could not be linked to prolonged cold storage of spores, nor to similar phenotypes reported
for a number of other mutants.
Furthermore, we developed a novel protocol for time-efficient isolation of Physcomitrella
antherozoids based on FACS of fluoresceín diacetate labelled antherozoids, a method crucial for future
studies of their methylation profiles.
Key-words: Physcomitrella patens; DNA methylation; de novo methyltransferases; DRM2
knockout; antherozoids; Fluorescence-activated cell sorting.
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Table of Contents
Acknowledgements ................................................................................................................................ i
Agradecimentos .................................................................................................................................... ii
Abstract ................................................................................................................................................. iii
Table of Contents ................................................................................................................................. iv
List of Abbreviations ............................................................................................................................ vi
List of Tables ......................................................................................................................................... ii
List of Figures ........................................................................................................................................ ii
Introduction
Epigenetics and DNA methylation ............................................................................................. 1
The sequence context of cytosine methylation and DNA methyltransferases..................... 3
DNMT2 and ribonucleic acid (RNA) methylation 4
CG methylation by MET1 and DNMT1 proteins 4
Chromomethyltransferases (CMTs) 5
De novo DMTases and CHH methylation 5
Physcomitrella patens as a model organism ......................................................................... 10
Cytosine methylation and DNA methyltransferases in Physcomitrella patens .................. 13
Physcomitrella patens transcriptomic atlas and specific expression of DNA
methyltransferases ............................................................................................................ 16
Aims of this study ..................................................................................................................... 18
Materials and Methods
Physcomitrella patens maintenance and growth .................................................................. 19
Confirmation of DRM2 deletion in the mutant lines by PCR................................................. 20
Fertilization rate assessment ................................................................................................... 21
Sporophyte collection and spore sterilization ....................................................................... 21
DRM2 K.O. and WT spores germination and colony growth assays ................................... 21
Colony growth after cold storage of spores 21
Colony area measurement and analysis 22
Colonies dry weight and analysis 23
WT spore germination and colony growth under different pH conditions 23
pSP3b plasmid construction .................................................................................................... 24
Transformation of WT line with linearized pSP3b plasmid ................................................... 28
DNA preparation: digestion and precipitation 28
Plant transformation 29
v
Selection of stable mutant lines 30
Genotyping of potential DNMT3b K.O. lines by multiplex in-tissue PCR ............................ 30
Antherozoids release and labeling assays ............................................................................. 31
Flow cytometry, cell sorting and microscopic confirmation of antherozoid samples ...... 32
Phylogenetic analysis of P. patens’ de novo DNA methyltransferases genes ................... 32
pAT05 plasmid re-sequencing and mapping ......................................................................... 33
Results
Phylogenetic analysis of Physcomitrella patens de novo methyltransferases .................. 34
Deep sequencing of pAT05 plasmid ....................................................................................... 35
Confirmation of DRM2 deletion lines Δdrm2#1 and Δdrm2#2 .............................................. 36
Differences in fertilization rate are only detected for Δdrm2#1 in the F0 generation ......... 39
Colonies appearance shows phenotypic variations after 21 days of growth ..................... 40
Δdrm2#2 colonies appear smaller and with a more regular shape than WT colonies when
germinated from spores stored at 4 ºC during 14 weeks. 40
Smaller colonies can be detected both in WT and Δdrm2 lines and do not seem to correlate
with the time of cold storage of spores 42
Colony growth phenotypes are not affected significantly by pH 43
Growth of P. patens colonies can be followed in detail by determination of colony area
and dry weight ................................................................................................................... 43
Determination of WT and Δdrm2 colonies’ area and its variation with the time of cold storage
of sterilized spores. 43
Cold storage of spores has little effect on dry weight of WT and Δdrm2 colonies. 48
Δdnmt3b knockout lines were not obtained ........................................................................... 50
Antherozoids lack autofluorescence and can be labelled with FDA ................................... 53
FACS sorting of antherozoids ................................................................................................. 56
Discussion ........................................................................................................................................... 62
Future Perspectives ............................................................................................................................ 69
References ........................................................................................................................................... 70
Supplemental Material
Tables ....................................................................................................................................... - 1 -
Figures ................................................................................................................................... - 13 -
vi
List of Abbreviations
5-mC: cytosine methylation
A. thaliana: Arabidopsis thaliana
AGO 4 / 6: Argonaute 4 / 6
BR: Broad range
CMT: Chromomethyltransferase protein
DCL3: Dicer-like 3 protein
DNA: deoxyribonucleic acid
DNMT1: (cytosine-5)-methyltransferase 1
DNMT2: DNA methyltransferase 2
DNMT3a/DNMT3b/DNMT3L: animal de
novo methyltransferase protein family
DMTase: DNA methyltransferase
DRM: domains rearranged
methyltransferase
dsRNA: double-stranded RNA
FACS: fluorescence-activated cell sorting
FDA: fluoresceín diacetate
GFP: green fluorescent protein
HF: high-fidelity
HR: homologous recombination
HS: high sensitivity
ICF: immunodeficiency, centromere
instability and facial anomalies
K.O.: knockout
KNOPS: standard media used in P. patens
cultivation
KNOPS+GT+H: KNOPS media
supplemented with glucose, nitrogen source
and hygromycin B antibiotic
KNOPS+T: KNOPS media supplemented
with a nitrogen source
lncRNA: long non-coding RNA
Mbp: mega base pairs
mCherry: a sub-type of a RFP protein
mdlc: minimal dicer-like gene
MET: methyltransferase
NGS: next-generation sequencing
nt: nucleotides
P. patens: Physcomitrella patens
PCR: polymerase chain reaction
PEG: polyethyleneglycol
Pol IV: RNA polymerase IV
Pol V: RNA polymerase V
PVP-40: polyvinylpyrrolidone-40
RdDM: RNA-directed DNA methylation
RDR 2 / 6: RNA-dependent RNA polymerase 2/6
RFP: red fluorescent protein
RNA: ribonucleic acid
RNAseq: RNA sequencing
SCs: sperm cells
siRNAs small-interfering RNAs
ssRNA: single-stranded RNA
TEs: transposable elements
tRNA: transfer-RNA
tRNAasp: aspartic acid tRNA
TxRed: Texas Red
WT#1: wild-type line grown with Δdrm2#1
WT#2: wild-type line grown with Δdrm2#2
WT: wild-type line
Δdnmt3b: dnmt3b gene deletion lines
Δdrm2#1: drm2 gene deletion line number 1
Δdrm2#2: drm2 gene deletion line number 2
Δdrm2: drm2 gene deletion lines
ii
List of Tables
Table 1: DNA methyltransferase (DMTases) genes present in the P. patens genome .................. 16
Table 2: Presence and absence call for the expression of Physcomitrella patens DNA
methyltransferases genes ........................................................................................................... 17
Table 3: KNOPS media constitution with the nutrients supplied to support Physcomitrella patens
tissue growth. .............................................................................................................................. 19
Table 4: Sperm-nutritive solution composition ................................................................................. 31
List of Figures
Figure 1: Schematic alignment of A. thaliana’s and Human’s (Homo sapiens) DNA
methyltransferases (DMTases) ..................................................................................................... 4
Figure 2: Model for RNA-directed DNA methylation (RdDM) canonical pathway, highlighting some
of the cellular players involved. ..................................................................................................... 7
Figure 3: Proposed mechanism for the inherence of cytosine methylation patterns across cell
divisions. ........................................................................................................................................ 8
Figure 4: Phylogenetic position of the major lineages of green plants............................................ 10
Figure 5: Physcomitrella patens life cycle. ...................................................................................... 11
Figure 6: P. patens sexual organs................................................................................................... 12
Figure 7: Cytosine methylation distribution across Physcomitrella patens genome.. ..................... 14
Figure 8: Process of the measurement of the colony’s area in ImageJ software ........................... 23
Figure 9: pSP3b plasmid map (with a total of 8256 nt). .................................................................. 27
Figure 10: Phylogenetic trees obtained from the analysis of the total set of DNA methyltransferases
sequences used in this work ....................................................................................................... 34
Figure 11: Map of the complete sequence of the pAT05 (with a total of 8201 nt), used to obtain
Δdrm2 lines. ................................................................................................................................ 36
Figure 12: Scheme of the approach used for the multiplex PCR to confirm the deletion of the DRM2
gene in Physcomitrella patens’ Δdrm2#1 and Δdrm2#2 line used in this work. ......................... 37
iii
Figure 13: Picture of the 1% agarose gel loaded with PCR products for wild-type (WT) and DRM2
mutant lines (Δdrm2#1 and Δdrm2#2) ........................................................................................ 38
Figure 14: Fertilization rates from wild-type (WT) grown with line 1 (WT#1) and line 2 (WT#2) as well
as for DRM2 mutant lines (Δdrm2#1 and Δdrm2#2). .................................................................. 39
Figure 15: F0’s wild-type colonies obtained 21 days after germination of spores ........................... 40
Figure 16: F0’s Δdrm2#2 colonies obtained 21 days after germination of spores. ......................... 40
Figure 17: F0 wild-type colonies obtained 21 days after germination of spores kept at 4 ºC for 3.5
months. ........................................................................................................................................ 41
Figure 18: Colonies from F0 Δdrm2#2 obtained 21 days after germination of spores, kept sterilized
at 4 ºC for 3.5 months ................................................................................................................. 41
Figure 19: Colonies of F0 wild-type (WT), obtained from the germination of spores stored at 4 ºC
during 6 weeks and of F0 Δdrm2#1 line colonies germinated from pores stores at 4 ºC for 10
weeks, after 21 days of growth. .................................................................................................. 42
Figure 20: Colonies from F0 wild-type germinated with water at different pH values, after 21 days of
growth. ......................................................................................................................................... 43
Figure 21: Variation of average colony area with different days of growth from WT#1 and Δdrm2#1
colonies, germinated from spores stored (0 to 14 weeks) at 4 ºC. ............................................. 45
Figure 22: Variation of the average colony area with the different days of growth from WT#2 and
Δdrm2#2 colonies, germinated from spores stored (0 to 14 weeks) at 4 ºC. ............................. 47
Figure 23: Scatter plot of the dry weight of the colonies of WT#1 and Δdrm2#1 lines after 21 days of
growth, obtained from the germination of spores stored at 4 ºC for different periods of time (0 to
14 weeks). ................................................................................................................................... 49
Figure 24: Scatter plot of the dry weight of the WT#2 and Δdrm2#2 colonies after 21 days of growth,
obtained from the germination of sterilized spores stored at 4 ºC for different periods of time (0 to
14 weeks). ................................................................................................................................... 50
Figure 25: Colonies obtained after two rounds of selection, regenerated from protoplasts subjected
to the transformation protocol with the pSP3b plasmid. .............................................................. 51
Figure 26: Scheme of the multiplex PCR approach used to genotype the selection-surviving colonies
of Physcomitrella patens transformed with the pSP3b plasmid. ................................................. 51
Figure 27: 1% agarose gel loaded with the in-tissue multiplex PCR reactions C and D, used for
genotyping the selection-surviving colonies from the transformation of Physcomitrella patens
protoplasts with the pSP3b plasmid ............................................................................................ 53
Figure 28: Bright field pictures of a single antherozoid and clusters of antherozoids ..................... 54
Figure 29: FDA labelled antherozoid ............................................................................................... 55
Figure 30: Labelled antherozoid clusters from Physcomitrella patens............................................ 55
iv
Figure 31: Isolated antherozoids obtained after antheridia sample filtering ................................... 56
Figure 32: Flow cytometric analysis of the flow through obtained using a 10 µm mesh. ................ 58
Figure 33: Flow cytometric analysis of the 28 µm mesh filtered flow-through sample.................... 59
Figure 34: Flow cytometric analysis of the 28 µm mesh filtered samples' flow-through ................. 60
Figure 35: Antherozoids sorted by FACS. ....................................................................................... 61
Figure 36: Examples of smaller, regular and round colonies lacking gametophores as reported for
some P. patens mutant lines ....................................................................................................... 67
1
Introduction
Epigenetics and DNA methylation
Epigenetics refers to a set of molecular mechanisms that affect the phenotype of a cell/organism
without affecting its genotype. All of these deoxyribonucleic acid (DNA) modifications compose the so
called Epigenome. It includes a range of chromatin modifications as DNA methylation, histone
modifications and positioning of nucleosomes (Feng et al., 2010a; Sasaki and Matsui, 2008). The most
common eukaryotic DNA modification is methylation of cytosine at position 5 of the pyrimidine ring
(Pélissier et al., 1999).
Cytosine methylation (5-mC) is present in most genomes of animals, plants, fungi, algae, protista
and bacteria (Noy-Malka et al., 2014). DNA methylation is a stable epigenetic modification in which an
enzyme known as DNA methyltransferase (DMTase) catalyzes the transfer of a methyl group from S-
adenosyl-L-methionine to the fifth carbon of a cytosine nucleotide in the DNA. This methylation of DNA
occurs after DNA synthesis and can be either maintained after cell division (in the case of symmetrical
sequence contexts, due to the presence of a 5-mC in the template strand) or deposited de novo (addition
of methyl groups to a target sequence devoid of pre-existing methylation) (Finnegan and Kovac, 2000;
Laird and Jaenisch, 1996).
DNA methylation has been proposed to function as a genomic immune system, in which invading
transposable elements (TEs - DNA regions capable of jumping in the genome) are recognized by the
host and methylated, suppressing their transcription in order to prevent further replication and preserving
a genome’s integrity from new mutations by insertions (Zemach and Zilberman, 2010). In both plants
and animals, 5-mC can be found in all regions of a genome and it constitutes an additional layer of
information that is known to be involved in the regulation of gene expression patterns (Law and
Jacobsen, 2011; Pélissier et al., 1999). Gene body methylation is present in diverse eukaryotic genomes
and is positively correlated with gene expression levels as opposed to methylation of TEs and other
repressive epigenetic marks that aim to silence these elements (Lister et al., 2009; Feng et al., 2010a;
Zemach and Zilberman, 2010; To et al., 2015).
In animals, between 3 to 8 % of all cytosines are known to be methylated and this occurs almost
exclusively in the symmetrical CG context, although some asymmetric methylated cytosines (CHH
sequence context, where H represents either nucleotide A, T or C), have also been observed in
embryonic stem cells, induced pluripotent stem cells, oocytes, male germ cells and in the brain
(Ichiyanagi et al., 2013). 5-mC residues are considered to be mutagenic and to increase C/G to T/A
mutation rates through spontaneous deamination that can be related to oncogenic point mutations (Laird
and Jaenisch, 1996). Aberrant de novo methylation is associated with silencing of tumor suppressor
genes in human cancers (Cao and Jacobsen, 2002b). In plant genomes, from 6 to 30 % of all the
cytosine nucleotides are known to be methylated and these are present in all three sequence contexts:
CG, CHG (symmetrical methylations contexts) or CHH (asymmetrical methylation) and most methylated
nucleotides occur within repetitive DNA found in heterochromatic regions (repeat rich and silenced
regions of the genome) (Finnegan and Kovac, 2000; Malik et al., 2012; Noy-Malka et al., 2014). Among
2
Arabidopsis thaliana's methylated residues approximately 24 % are GC, 6.7 % CHG and 1.7 % are CHH
methylated (Zhang et al., 2013).
Besides being implicated in the regulation of gene expression patterns, DNA methylation also
affects cellular differentiation, developmental programs and genome stability. While loss of genome
methylation is lethal in vertebrate embryos, plants are able to tolerate and survive, although pleiotropic
defects may be observed (Malik et al., 2012). Epigenetic gene silencing is therefore important in
developmental phenomena such as imprinting (gene expression in a parent-of-origin manner) in both
plants and mammals, as well as in cell differentiation and reprogramming in order to maintain genome
integrity from generation to generation. To this end TEs and repetitive DNA elements must be kept under
tight regulation in reproductive cells, to avoid mutations to be transmitted to the next generation (Feng
et al., 2010a; Finnegan and Kovac, 2000).
Epigenetic reprogramming is a process wherein global changes in the epigenome take place and
it occurs both in plants and animals, although with some differences. It involves both DNA demethylation
(removal of 5-mC from the DNA) and remodeling of histones on a genome-wide scale in the germ line
of these organisms, followed by the re-setting of epigenetic marks in the early embryo (Feng et al.,
2010a; Kawashima and Berger, 2014; Morgan et al., 2005; Sasaki and Matsui, 2008) In plants some
genetic elements, such as TEs, seem to be persistently silenced by DNA methylation across
generations, leading to the idea that DNA methylation might be inherited in a stable manner from one
generation to the next and that epigenetic reprogramming might not exist in plants. However, genome-
wide analyses of the epigenome detected an overall reduction of DNA methylation in the plant germ line
during gametogenesis and showed that epigenetic reprogramming takes place during plant sexual
reproduction (Calarco and Martienssen, 2011; Jullien et al., 2012).
Today, it is well accepted that DNA methylation patterns are dynamic during plant development.
Genome-wide losses of DNA methylation are known to occur during both male and female
gametogenesis followed by de novo methylation after fertilization (Law and Jacobsen, 2011). As in
mammals, and in order to maintain genome integrity from generation to generation, TEs and repetitive
DNA elements must also be tightly regulated in reproductive cells of plants. One mechanism used to
achieve this is through the stable inheritance of DNA methylation in TE-rich regions. Although, plants
are not known to undergo genome-wide waves of demethylation in their germ line as mammals do, most
of plant's reprogramming occurs in non-germ line reproductive cells and it may function to reinforce
silencing of TEs in the germ cells (Feng et al., 2010a; Slotkin et al., 2009).
After fertilization, the somewhat demethylated genome of the plant’s embryo seems to undergo
de novo DNA methylation during embryogenesis, which is mediated by de novo DMTases, as in
mammals (Kawashima and Berger, 2014). Likewise, in both groups of organisms a genome-wide
demethylation occurs in the extra embryonic tissues (endosperm and placenta) but not in the embryo,
which indicates that epigenetic regulation in the extra embryonic tissues is different from the embryonic
one, being conserved (or reinvented) in plants and animals. In both cases variations in DNA methylation
levels between somatic cells and gametes involves loss of total DNA methylation but gains of CHH
methylation through de novo DNA methylation (Feng et al., 2010a; Zhang et al., 2013).
3
As DNA methylation, DNA demethylation is also dynamic and the equilibrium between these
processes is essential for cell survival. Therefore the action of DMTases enzymes is balanced with the
action of DNA demethylating glycosylase enzymes, such as Demeter and Repressor of Silencing 1.
However, passive demethylation is also known to occur by deamination and base excision repair
processes, and these effects are responsible for the variable nature of the methylome (the set of all the
methylated residues of a genome) (Chen et al., 2012; Ooi and Bestor, 2008; Zheng et al., 2008).
The sequence context of cytosine methylation and DNA methyltransferases
DNA methylation is achieved either by methylating a cytosine residue de novo or by maintaining
a previously established pattern of 5-mC residues (Malik et al., 2012). Cytosine methyltransferase
enzymes (DMTases) catalyze the transfer of an activated methyl group from S-adenosyl methionine to
the 5 position of the cytosine ring. Arabidopsis thaliana (A. thaliana) has at least 10 genes that could
encode DMTases (Martienssen and Colot, 2001). As explained above, DNA methylation occurs in three
different sequence contexts in plant genomes: CG, CHG - symmetrical contexts; and CHH (H = A, C or
T) - asymmetrical context (Feng et al., 2010a).
After each round of DNA replication in cell division, each daughter cell carries hemimethylated
DNA (one methylated parental strand and one newly synthesized unmethylated strand). For DNA
sequences methylated in the CG and CHG context, the methylation can be established on the
unmethylated strand by maintenance DMTases based on the information from the parental methylated
strand, following a semi-conservative mechanism (Cao and Jacobsen, 2002b; Cao et al., 2000;
Mahfouz, 2010). Methylation that occurs at previously unmethylated cytosines is de novo methylation.
For symmetric sites, de novo methylation needs to occur only once, after which methylation can be
preserved by maintenance activity, however for methylation at asymmetric sites (CHH context) de novo
methylation must occur continuously (Cao and Jacobsen, 2002b; Cao et al., 2000; Zhang et al., 2013).
Therefore, the pattern of cytosine methylation is established by de novo DMTases and maintained
by maintenance methyltransferase activities. In plants, the DMTases are categorized into four
subfamilies: DNA methyltransferase 2 (DNMT2), methyltransferases (METs),
chromomethyltransferases (CMTs) and domains rearranged methyltransferase (DRMs) (Malik et al.,
2012). In animals, MET proteins are known homologs of the (cytosine-5)-methyltransferase 1 (DNMT1)
proteins and DRM proteins are replaced by the de novo methyltransferases group constituted by DNMT3
proteins. While DNMT2 protein family is conserved between both groups, CMT family is specific to plants
(Kuhlmann et al., 2014).
Different DMTase families are thought to be responsible for cytosine methylation in different
sequence contexts (Kuhlmann et al., 2014). In Figure 1, a schematic alignment between A. thaliana's
and Human DMTases can be seen. It is thought that members of MET, CMT and DNMT subfamilies
originated from a common ancestral gene that possibly also gave rise to the plant and animal de novo
DMTases, while the lineage that gave rise to present day DRMs appears to have diverged earlier than
the bifurcation of plant and animal lineages (Malik et al., 2012).
4
Figure 1: Schematic alignment of A. thaliana’s and Human’s (Homo sapiens) DNA methyltransferases
(DMTases). The similarities between the domains from the same family are clear: A. thaliana’s MET1 and H.
sapiens DNMT1 are very similar, the major difference is the zinc finger domain present in DNMT1 and absent in MET1; DRM2 protein is specific to plants but similar to human DNMT3a and DNMT3b involved in CHH methylation. CMT protein is plant specific, sharing the cytosine methyltransferase domain with the other proteins but having a chromo domain and a BAH domain, absent in all other DMTases (Henderson and Jacobsen, 2007).
DNMT2 and ribonucleic acid (RNA) methylation
Plants, animals and fungi share the highly conserved DNMT2 proteins, that contain all catalytic
motifs expected of a DMTase, but show no such activity in vitro (Zemach and Zilberman, 2010). Cytosine
DMTases belonging to DNMT2 subfamily are among the most well conserved in plants (Malik et al.,
2012), but they do not appear to play a significant role in establishing or maintaining DNA methylation
patterns. Besides not showing DMTase activity in vitro, DNMT2 loss of function mutations do not show
any reduction in the amount of DNA methylation (Cao et al., 2000).
While the possibility that DNMT2 can function as a DMTase remains, evidence that DNMT2 is a
specific transfer-RNA (tRNA) methyltransferase with conserved functions across mammals, flowering
plants and insects comes from a study conducted by Goll and co-workers (2006) where purified DNMT2
from human cells was able to methylate specifically the aspartic acid tRNA from human, mouse,
Drosophila melanogaster and A. thaliana samples (Goll et al., 2006).
CG methylation by MET1 and DNMT1 proteins
In both mammals and plants, symmetrical CG methylation is maintained by the maintenance DNA
methyltransferase termed DNA (cytosine-5)-methyltransferase 1 (DNMT1) in mammals or DNA
methyltransferase 1 (MET1) in plants (Feng et al., 2010b). These two proteins are very similar in both
sequence and function and in Arabidopsis, loss-of-function MET1 mutants and antisense-met1
transgenic plants, lack the majority of CG methylation (Cao and Jacobsen, 2002b).
Repetitive sequences are densely methylated in all sequence contexts. In A. thaliana’s MET1
mutant, loss of CG methylation seems to be enough to reactivate transposons, which may indicate a
role for CG methylation in silencing TEs. AtMET1 mutant also exhibits morphological defects such as
delayed flowering and reduced size (Mahfouz, 2010; Noy-Malka et al., 2014).
Since, DNMT1 has a catalytic preference for hemimethylated substrates, this provides an
attractive model for the efficient maintenance of CG methylation after DNA replication and during cell
5
division, since the parental DNA strand should be methylated and this may be the signal for the
methylation of the symmetrical cytosine in the newly synthesized DNA strand (Finnegan and Kovac,
2000; Henderson and Jacobsen, 2007).
Chromomethyltransferases (CMTs)
The CMT DMTase family is found exclusively in plants. These DMTases are characterized by the
presence of a chromatin organization modifier (chromo) domain in its C-terminal region and a bromo
adjacent homology (BAH) domain in the N–terminal regulatory region (Figure 1) (Cao et al., 2000;
Henikoff and Comai, 1998; Mahfouz, 2010).
There are three related CMT genes in A. thaliana and Oryza sativa (rice). In A. thaliana, while
CMT1 is predicted to be non-functional, being truncated by a transposon insertion, CMT2 and CMT3
have been shown to be functional (Henikoff and Comai, 1998; Martienssen and Colot, 2001; Zemach et
al., 2013). In most plant species, only CMT3 and its homologs are known to be actively transcribed and
have been functionally characterized (Henikoff and Comai, 1998).
The maintenance of DNA methylation in symmetric CHG context is unique to plants, and in A.
thaliana the majority of CHG context methylation depends on CMT3 (Kuhlmann et al., 2014; Zhong et
al., 2014). CMT3 loss-of-function mutants were isolated in three independent studies, showing a
genome-wide loss of CHG methylation and a reduction on CHH methylation at some loci. These mutants
do not display any morphological abnormalities and only the triple mutant, CMT3 DRM1 DRM2, shows
pleiotropic developmental defects such as partial sterility and reduced plant size, indicating some
overlap in CMT3 and DRMs function in non-CG methylation (Cao and Jacobsen, 2002a, 2002b). In
maize loss of function of CMT3’s homolog ZMET2 shows reduced CHG methylation at centromeric
repeats but no changes in other sequence contexts (Papa et al., 2001).
Arabidopsis CMT2 diverges from CMT3 in the N–terminal regulatory region and is unable to
complement loss-of-function of CMT3 mutants’ changes on CHG distribution (Henikoff and Comai,
1998). In 2013 Stroud et al., showed that CMT2 can methylate both CHG and CHH sites, but since
CMT2 mutants show a global reduction of CHH methylation while CHG methylation remains mostly
unaffected, CMT2 shows different sequence specificities than CMT3 by preferentially methylating CHH
sites independently of the action of DRM2 (Stroud et al., 2013; Zemach et al., 2013).
Therefore, the collective activity of CMT3, CMT2 and DRMs is responsible for all non-CG
methylation detected in A. thaliana’s genome (Stroud et al., 2013).
De novo DMTases and CHH methylation
Asymmetric methylation (CHH) is maintained by the persistent activity of de novo
methyltransferases capable of methylating previously unmethylated DNA. CHH methylation is abundant
in plants and it can be maintained either by the RNA-directed DNA methylation (RdDM) pathway that
requires DRMs action, or independently of this process by CMT2’s activity (described above) (Cao and
Jacobsen, 2002b; Cao et al., 2000; Stroud et al., 2013; Zhang et al., 2013).
6
CHH methylation is also present at detectable levels in mammals, especially in stem cells, and
this methylation is introduced by de novo DMTases: DNMT3a and DNMT3b, that are also required for
the maintenance of CG methylation at some loci (Feng et al., 2010b).
Both DRM1 and DRM2 contain catalytic domains showing sequence similarity to those of the
mammalian DNMT3 methyltransferases. However, unlike DNMT3s, the DRMs have unique N-termini
containing ubiquitin associated domains (Cao and Jacobsen, 2002b) and a different order of catalytic
motifs, reason why the A. thaliana's proteins have been named the domains rearranged
methyltransferases (DRMs) (Cao et al., 2000).
The domains rearranged methyltransferases (DRMs) in plants
DRM genes were reported by Cao et al., 2000 as genes required for de novo DNA methylation in
A. thaliana since drm1 drm2 double mutants lacked the de novo methylation normally associated with
transgene silencing (Cao et al., 2000). Therefore DRMs are key de novo methyltransferase in plants,
but how they act mechanistically is poorly understood (Zhong et al., 2014).
A. thaliana DRM1 DRM2 double mutants revealed no morphological defects although they display
subtle changes in the methylation patterning (Mahfouz, 2010) however, DRM1 DRM2 CMT3 triple
mutants show developmental phenotypes which include misshapen leaves and reduced stature
(Henderson and Jacobsen, 2007). According to Cao et al., (2003), neither DRM nor CMT3 mutants
affected the maintenance of pre-established CG methylation. However, DRM mutants showed a nearly
complete loss of asymmetric methylation and a partial loss of CHG methylation. Furthermore, RdDM
requires the activity of DRM2 to establish methylation in all sequence contexts (Kuhlmann et al., 2014;
Naumann et al., 2011). DRM1 single mutants show no particular defects or alterations on the
methylation pattern and are not considered to be required for methylation in any sequence context while
DRM3 is required for full levels of DRM2 mediated DNA methylation, being now considered a weak
factor for the RdDM pathway (Zhong et al., 2014) that will be briefly described below.
The RNA-directed DNA methylation pathway (RdDM)
RNA-directed DNA methylation (RdDM) is a small RNA-mediated epigenetic modification that so
far was only detected in plants. It leads to cytosine methylation of the DNA region complementary to the
RNA sequence (Aufsatz et al., 2002; Law and Jacobsen, 2011; Naumann et al., 2011).
The RdDM pathway (Reviewed in Matzke et al. 2014 and Movahedi et al. 2015) involves two main
stages (Figure 2):
First, the phase wherein small-interfering RNAs (siRNAs) are synthetized. This stage starts with
a plant-specific RNA polymerase IV (Pol IV) generating single-stranded RNA (ssRNA) transcripts, which
then are copied into double-stranded RNA (dsRNA) by RNA-dependent RNA polymerase 2 (RDR2).
The resulting dsRNA molecules are cleaved in 23-24 nt (nucleotides) siRNA by dicer-like endonuclease
3 (DCL3) and loaded into Argonaute 4 (AGO4) forming AGO4-siRNA complexes (Figure 2, 1st phase)
(Wassenegger et al., 1994).
7
Figure 2: Model for RNA-directed DNA methylation (RdDM) canonical pathway, highlighting some of the
cellular players involved. Despite the usual RNAi players, as dicer-like proteins (DCL) and argonaute proteins (AGO), some other players are involved in RdDM, such as: RDRs (RNA-dependent-RNA polymerases), two specific plant polymerases (Pol IV and Pol V), the de novo methyltransferase DRM2, and other proteins known to be involved in this process (e.g. SHH1 and RDM1). The process starts with the 1st phase, known as the biogenesis of siRNA. This starts with the transcription of non-coding regions by Pol IV afterwards, RDR2 will copy this transcript making double-stranded RNA (dsRNA) which is then cleaved to 24 nt (nucleotides) siRNA by DCL3. These siRNAs are then loaded into AGO/RISC (RNA-induced silencing complex). The process proceeds to the 2nd phase, known as the siRNA targeting and methylation stage, wherein AGO/RISC loaded siRNA are recruited to the target loci due to interactions of the complex with Pol V and its transcripts, followed by the recruitment of DRM2 de novo methyltransferase that will deposit methyl groups in the cytosine residues of the DNA region correspondent to the siRNA. Adapted from (c).
Second, the targeting and methylation phase: this involves another plant-specific RNA
polymerase, polymerase V (Pol V), which produces long-non-coding RNA (lncRNA) transcripts that are
proposed to act as a scaffold to recruit AGO4 through base-pairing of associated siRNAs since Pol V's
largest subunit -NRPE1.7- interacts with AGO4 to recruit it together with the bound siRNAs (Böhmdorfer
et al., 2014; Law and Jacobsen, 2011; Zheng et al., 2009; Zhong et al., 2014). Then the siRNAs, AGO4
and its associated proteins are recruited to the siRNA's correspondent DNA sequence and facilitate
target methylation by the guidance of DRM2 de novo methyltransferase to that specific locus
(Böhmdorfer et al., 2014) (Figure 2, 2nd phase). The machinery by which siRNAs target cytosine
methylation to the correspondent DNA sequence is poorly understood and could involve either DNA–
RNA or RNA–RNA hybridization events (Henderson and Jacobsen, 2007).
Various biological roles for RdDM have been proposed, such as the silencing of TEs present in
eukaryotic genomes (to avoid transposition and genome damage), maintenance of chromatin structure,
gene imprinting, gene regulation, plant development and stress responses (Böhmdorfer et al., 2014;
Viswanathan and Jian-Kang, 2011).
More recently, long and centromeric transcriptionally active TEs were shown to be methylated by
a non-canonical RdDM pathway that involves RNA polymerase II dependent transcripts. These are
8
converted to dsRNA by RDR6, processed into 21-22 nt siRNAs and associate with AGO6 to guide de
novo methylation. This RDR6-RdDM pathway is particularly active in the precursor cells of the
reproductive tissues (young flower buds), where AGO6 is expressed. Besides, RDR6-RdDM functions
independently of Pol IV-RdDM although they can complement each other to fully silence active TEs
(Eamens et al., 2008; Garcia et al., 2012; Mccue et al., 2015; Pontier et al., 2012; Stroud et al., 2013).
In summary, RNA-directed DNA Methylation
(RdDM) establishes de novo methylation in all sequence
contexts by the action of DRM2 (Figure 3). Maintenance
of the methylation can be achieved by MET1 and CMT3
for symmetrical sites (Figure 3, loop 1) and CHH
methylation patterns can be inherited across cell divisions
by continuous action of RdDM (Figure 3, loop 2) or CMT2
independently of RdDM (Bond and Baulcombe, 2014; Cao
and Jacobsen, 2002a; Cao et al., 2000; Zemach et al.,
2013; Zhang et al., 2013).
Figure 3: Proposed mechanism for the inherence of cytosine methylation patterns across cell divisions. Methylation of previously unmethylated regions occurs by RNA-directed DNA methylation (RdDM), targeted by small-RNAs (sRNA) and by the action of domains rearrangement methyltransferase (DRM2). Maintenance of the previously established pattern of methylation can be achieved either by a sRNA-independent mechanism – loop 1, wherein MET1 and CMT3 enzymes maintain symmetrical methylation patterns or by continuous sRNA targeting in the RdDM pathway for the CHH methylation context – loop 2 (Bond and Baulcombe, 2014).
DNMT3
The first evidences of DNMT3a and DNMT3b activity as DNA methyltransferases in vivo comes
from a study conducted in 1999 by Hsieh, this same study also suggested that these enzymes could
have different target requirements due to their differential occupancy in the nucleus.
Okano et al., (1999) found that DNMT3a and DNMT3b are essential for de genome-wide de novo
methylation and for mammalian development since both DNMT3a and DNMT3b heterozygous mice
were normal and fertile, however most homozygous mutant mice died during embryogenesis, showing
that DNMT3a and DNMT3b have overlapping functions during early embryogenesis. In the same year,
Xie et al. showed that DNMT3a encoded a polypeptide with 912 amino acid residues that was ubiquitous
expressed in most mice adult tissues, whereas DNMT3b with 853 amino acids was detected at lower
levels in adult tissues except on testis, thyroid and bone marrow where it was highly expressed.
Moreover both genes were overexpressed in most tumor cell lines studied (Xie et al., 1999).
The recessive autosomal disorder known as ICF syndrome (immunodeficiency, centromere
instability and facial anomalies) is characterized by a variable immunodeficiency, mild facial anomalies,
demethylation of satellite repeats and centromeric decondensation that leads to chromosomal instability,
causing most ICF patients to succumb to infectious diseases before adulthood. It was the first human
genetic syndrome to be associated with defects in methylation patterns, being caused by mutations on
both alleles of DNMT3b gene (Hansen et al., 1999; Xu et al., 1999), that is considered to be essential
for the methylation of the satellite repeats and centromeric repeats (Kato et al., 2007). DNMT3b
9
polymorphisms were also linked to progression of joint destruction in rheumatoid arthritis (Nam et al.,
2010) and to the occurrence of Down’s syndrome in children due to a failure of normal chromosomal
segregation during meiosis (that is considered to be the origin of 90% of all Down’s syndrome cases)
(Coppedè et al., 2013; Jaiswal et al., 2015).
Further studies on these enzymes revealed that the DNMT3a and DNMT3b proteins are
expressed at different stages of embryogenesis, suggesting their involvement on the selective de novo
methylation in the inner cell mass (cells that will originate the new organism) (Rhee et al., 2002;
Watanabe et al., 2002), but with distinct functions, since DNMT3b showed a higher activity in non-CG
methylation than DNMT3a (Suetake et al., 2003). Both proteins appear to be critical to regulate the
Immunoglobulin kappa light chain rearrangement during the early development of B-lymphocytes in
humans (Manoharan et al., 2015) and DNMT3b was also demonstrated to cooperate with DNMT1 in
order to maintain DNA methylation and tumour suppressor gene silencing in human cancer cells (Rhee
et al., 2002). In 2012, Chen and co-workers showed that DNMT3a and DNMT3b (but not DNMT1) could
also act as DNA dehydroxymethylases (enzymes capable of converting 5-hydroxymethyl cytosine, an
oxidized form of 5-mC, directly to cytosine) and therefore participate in DNA demethylation, depending
on the redox environment of the cell.
In animals, in addition to DNMT3a and DNMT3b, the DNMT3 family includes an enzymatically
inactive paralogue: DNMT3L, a regulatory factor that complexes with DNMT3a and/or DNMT3b in order
to stimulate their activities (Sasaki and Matsui, 2008), being not only required for normal male meiosis
in mice, for the heritable silencing of retrotransposons in male germ cells (Bourc’his and Bestor, 2004)
but, together with DNMT3a, also for the de novo DNA methylation of imprinted genes in mammalian
germ cells (Jia et al., 2007). Recently, it was shown that DNMT3L functions in vivo by regulating CG
versus non-CG substrate preference of DNMT3A and DNMT3B (Tiedemann et al., 2014).
Depletion of all DNMT3 family members results in the hypomethylation of almost all non-CG sites,
confirming that these enzymes are responsible for the non-CG methylation in mammals (Tiedemann et
al., 2014) and their different functions and specificities were explored in more detail recently. Auclair et
al., (2014) showed that the onset of genome-wide methylation in the early epiblast is correlated with the
upregulation of DNMT3a and DNMT3b genes, with DNMT3b’s mRNAs reaching higher levels of
expression. They also verified that the inactivation of one DNMT3 gene does not modify the expression
of the other DNMT genes in mouse embryos and that the inactivation of either DNMT3a or DNMT3b
leads to a partial reduction in global methylation, indicating that the inactivation of one enzyme is
compensated by the other and that both enzymes cooperate to methylate the bulk of the genome.
Overall, the inactivation of DNMT3b leads to a higher number of hypomethylated sequences indicating
that DNMT3b has a greater contribution to methylation of the mammalian genome than DNMT3a
(Auclair et al., 2014).
In human oocytes DNMT3b showed a 10-fold higher expression than DNMT3a, and DNMT3L
was not expressed, suggesting that DNMT3b may be the critical de novo DNA methyltransferase during
human oocyte development (Okae et al., 2014). DNMT3b is known to be required for the de novo
methylation of particular regions of the genome. Mutations on this enzyme in human and mice can lead
to deficient methylation of pericentromeric repetitive DNA sequences and CpG islands (regions rich in
10
CG methylation), or to X chromosome inactivation. These alterations suggest that DNMT3b can modify
regions of already silenced chromatin (Auclair et al., 2014; Bird, 2002; Tiedemann et al., 2014).
Physcomitrella patens as a model organism
With a genome size of approximately 480 Mbp (mega base pairs) distributed across its 27
chromosomes, the model moss Physcomitrella patens (P. patens) represents the first bryophyte to have
its genome sequenced. Bryophytes comprise hornworts, mosses, and liverworts and were the first plants
to colonize land (Rensing et al., 2008). It is estimated that bryophytes and flowering plants evolution
diverged between 450 and 500 million years ago, a similar evolutionary distance as observed between
humans and fishes (Arif et al., 2013; Mosquna et al., 2009). More than 30 % of the assembled gene
products have detectable homologues in seed plants and more than 66 % of A. thaliana's genes have
detectable homologs in P. patens.
Figure 4: Phylogenetic position of the major lineages of green plants. Mosses (such as Physcomitrella
patens), liverworts and hornworts compose the bryophytes. More ancestral groups (below liverworts) comprise algae, and more evolved groups (above hornworts) comprise lichens (lycophytes), monilophytes and finally more recent plants belonging to gymnosperm and angiosperm groups. The nodes represent landmarks for evolution, with new characteristics that emerged indicated by arrows. Characteristics with asterisks (*) may have evolved after the point indicated by independent processes. The designation for the groups is written in bolt black. Species belonging to each group, that have their genome sequenced are written in blue, if their genome sequence is expected in a near future the species are between parenthesis (Prigge and Bezanilla, 2010).
These characteristics of P. patens’ genome, together with its ideal phylogenetic position, makes
it a good model for developmental studies and to unravel the evolutionary changes that allowed the
conquest of land by plants (Rensing et al., 2008; Nishiyama et al., 2003).
Today, all land plants are characterized by an alternation of two generations: the haploid
gametophyte and the diploid sporophyte generations. In flowering plants, the sporophyte comprises
complex organs including leafy shoots and flowers, while the gametophyte is composed of their gametes
11
(the egg cell and the sperm cells - SCs) that only develop during the plant's adult phase. However in
bryophytes, the gametophytic generation is the photosynthetically active one and dominates over the
shorter sporophyte generation (Mosquna et al., 2009; Nishiyama et al., 2003). Another main difference
between flowering plants and bryophytes is that, while flowering plants propagate by diploid seeds,
bryophytes disseminate through haploid spores (Mosquna et al., 2009). Fossil evidence suggests that
early land plants were structurally similar to bryophytes given that they most likely had a dominant
haploid phase and were dependent on water for sexual reproduction by relying on motile male gametes
(Rensing et al., 2008).
Figure 5: Physcomitrella patens life cycle. Most of the P. patens’ tissues belong to the haploid generation and the short diploid phase is only obtained after fertilization and before meiosis. Haploid generation –
Gametophyte: starts with spore’s germination forming a branched filamentous protonema tissue that comprises the chloroplast enriched chloronema and the caulonema (protonema cells containing less chloroplasts). The formation of meristematic buds marks the shift from the juvenile protonema to the adult gametophyte that develops gametophores (leaf-like structures) and rhizoids at its base. The sexual reproduction phase involves the development of the female archegonium and male antheridium. Within antheridia, motile spermatozoids (or antherozoids, the male gametes) are formed. When released these will fertilize the single egg cell (female gamete) within the archegonium. Diploid generation – Sporophyte: after fertilization the diploid zygote is formed. This zygote will grow into a spore capsule where the spores will mature. Within the spore capsule meiosis takes place and about 4000 to 6000 haploid spores are produced. After ripening, the spore capsule releases the spores, whose germination will give rise to new organisms in the gametophytic generation (Strotbek et al., 2013).
In P. patens the gametophytic generation comprising the haploid tissues represents the
predominant generation. Its life cycle takes between 3 to 4 months under standard culture conditions
used all over the world, starting from the germination of a haploid spore that forms a branched
filamentous protonema tissue composed of two distinct cell types: the chloroplast enriched chloronema
cells and the fast-growing caulonema cells containing less chloroplasts (Figure 5) (Strotbek et al., 2013).
The success of spore germination is a pre-requisite for the establishment of a plant in a new
location. It starts with the swelling of the spore, due to the uptake of water, until the rupture of the cell
wall takes place leading to the formation of a germ tube. So far, all bryophytes were found to require
water of the germination of their spores (Glime, 1983). Freezing of the spores was found to be favorable
to the germination of some species, although only if it would occur after spore hydration. Spores are
also considered to be less susceptible to cold damage (During, 1979). In 1939, Apinis reported that
12
spores of most mosses could germinate in a wide range of pH values, although the growth of the
protonema could be affected. After germination, the growth of the protonema starts by the development
of chloronema cells. The transition from chloronema to caulonema growth occurs within the first 7 days
of growth, after high density of chloronema cells is achieved or nutrients are depleted (Jang and Dolan,
2011; Prigge et al., 2010).
The transition from the juvenile protonema tissue to the adult gametophyte is initiated by the
formation of meristematic buds with three apical cells that develop into gametophores with a leaf-like
shape and rhizoids at its base. Sexual reproduction is initiated by the development of both female
archegonia (Figure 6 A) and male antheridia (Figure 6 B), gamete producing organs at the tip of the
gametophore (Schaefer and Zrÿd, 2001; Strotbek et al., 2013). The development of P. patens sexual
organs was described in 2013 by Landberg et al., in this study the authors divided the development of
both organs in 10 stages, showing that organ morphogenesis is highly organized in P. patens. The first
organ to be formed is the male antheridium in the center of the primary shoot apex. Then the next
antheridia will start to develop flanking the first one, and the same occurs for the female organs.
As the male antheridium starts to develop, two different cell types start to emerge: the outer cell
layer that divides to keep up with the increasing size of the organ and the inner spermatogenous cells.
By the time that the inner cells acquire a round shape (stage 7), the outer cells produce a yellow pigment
and the most apical cells start to swell. Afterwards (stage 8), the inner cells – spermatids - have
undergone the final cell division and will start spermatogenesis until stage 9 is reached. In stage 9, the
sperm cells (SCs) or antherozoids are considered to be mature, biflagelated, slender and coiled cells
that will be released in stage 10 after the bursting of the swollen apical cells. In standard growth it takes
14 days since the start of the development of the antheridia to the release of the mature SCs (Landberg
et al., 2013). Physcomitrella patens’ antherozoids are strikingly similar to the sperms of certain animals
in the way that they are flagellated cells and swim. They are elongated and coiled cells that possess no
cell walls (Paolillo, 1981; Wolniak et al., 2000).
The female organ – archegonium – starts developing several days later than the first antheridium.
It is only at stage 5 that the egg cell precursor starts to enlarges and at stage 7 it divides asymmetrically
giving rise to a cell that will mature and form the egg cell (on stage 8)
and to a smaller upper cell, that will die during stage 8. It is also during
stage 7 that the archegonium’s neck is formed. At the 9th stage the egg
cell is fully matured (indicated by the red arrow in Figure 6 A), the canal
of the archegonium opens so that fertilization can take place. At stage
10 the outer neck cells lose their contents and die. The process of
development and maturation of the archegonia takes 9 days and the
first archegonium is ready to be fertilized at the same time as the first
antheridium bursts and releases the sperm (Landberg et al., 2013).
Figure 6: P. patens sexual organs. A: Drawing of a mature archegonium (female sex organ), within which the single egg cell (indicated by a red arrow) lays in its cavity; B: drawing of a mature antheridium releasing its antherozoids that will swim in water until they find and fertilize the egg cell, giving rise to the diploid zygote. Adapted from Reski, 1998.
13
This makes P. patens a monoecious species (meaning that an organism has both sexes) and
both male and female organs are found interspersed in the same gametangia. Since both male and
female reproductive organs are mature at the same time, organ maturation is not a barrier to self-
fertilization in P. patens (Landberg et al., 2013; Strotbek et al., 2013). Upon fertilization, the diploid
sporophyte (apical spore capsule) starts to develop from the zygote, supported by a short seta. Inside
this capsule, meiosis will take place and about 4000 to 6000 haploid spores are produced. These spores
will mature inside the sporophyte and afterwards, the capsule breaks open releasing the spores for
propagation (Figure 5) (Strotbek et al., 2013).
Many hormonal factors studied in angiosperms and absent from unicellular algae, such as auxins,
cytokinins and abscisic acid, are known to be present in P. patens (Rensing et al., 2008; Schumaker
and Dietrich, 1997) and to be involved in developmental transitions (e.g. caulonema differentiation
involves auxins and cytokinins) (Cove and Ashton, 1984). In addition, the perception of auxins was
already studied in P. patens and was found to be a conserved pathway among land plants, despite the
different roles played by these phytohormones (Prigge et al., 2010).
Almost any tissue or single cell of P. patens is able to regenerate into intact plants on hormone-
free media which makes its cultivation and vegetative propagation possible and easy. The ability for cell
cultivation in liquid media up to 100 L volumes, together with the high ability to perform post-translation
protein modifications make P. patens a good alternative for the production of bio-pharmaceuticals
(Prigge and Bezanilla, 2010; Strotbek et al., 2013). Besides this, P. patens is easy to handle and does
not require expensive material, reagents or much space to be maintained in a laboratory (Schaefer and
Zrÿd, 2001).
However, P. patens main advantage that may be the reason why it has emerged as a new plant
model is the established techniques to genetically modify it. Since P. patens shows a high frequency of
homologous recombination (HR) similar to that of Saccharomyces cerevisiae, it allows an efficient gene
targeting and the generation of stable mutant lines to conduct reverse genetics experiments and to
analyze gene function (Prigge and Bezanilla, 2010; Reski, 1998; Schaefer and Zrÿd, 2001; Strotbek et
al., 2013). Moreover, altering or deleting a gene in a haploid organism can potentially lead to an altered
phenotype, making forward and reverse genetic approaches more straightforward in P. patens than in
diploid seed plants such as Arabidopsis or rice, assuming that the gene’s mutation is not lethal when in
a haploid state (Reski, 1998).
Due to these main advantages, P. patens allows the study of gene function and enables the
decoding of developmental mechanisms present in ancestral land plants. By using evolutionary
approaches, it is also possible to identify mechanisms that represent innovations in flowering plants and
to understand the colonization of terrestrial environments by plants (Prigge and Bezanilla, 2010).
Cytosine methylation and DNA methyltransferases in Physcomitrella patens
In the recent years, evidences that mechanisms for gene silencing play a role in regulating
developmental programs have been increasing. As discussed above, gene silencing can involve nucleic
acid based mechanisms, such as DNA methylation and/or histone modifications (Feng et al., 2010b;
Mosquna et al., 2009; Sasaki and Matsui, 2008). It is known that DNA methylation is essential to regulate
14
important developmental processes in higher eukaryotes, but little is known about its necessity and role
in early land plants such as P. patens (Malik et al., 2012).
In 2009, Mosquna and co-workers reported the P. patens gene PpFIE as an ortholog of the
polycomb group complex, that is known to control gene expression epigenetically and to be involved in
the methylation of lysine 27 residue on histone 3, being in part responsible for gene silencing and
changes in developmental programs in A. thaliana. In 2010, Prigge and Bezanilla reported that the
polycomb repressive complex 2 was required to repress sporophyte development in the gametophyte
stem cell during the apogamy process in P. patens, wherein cells from the gametophyte other than the
egg cell initiate sporophyte development. This led to the idea that the transition of the different stages
of a plant's life cycle are likely to involve epigenetic mechanisms (Pires and Dolan, 2012).
In P. patens, all three contexts of cytosine methylation (CG, CHG and CHH) were found to be
enriched in repetitive regions (such as TEs), reduced on gene bodies and almost absent around the
transcriptional start site (Figure 7). The levels of 5-mC residues present on P. patens whole plants
nuclear genome were found to be around 29.5 % of CG, 29.7 % of CHG and 23.2 % of CHH methylated
cytosines (Zemach et al. 2010). P. patens levels of CHH methylation (~ 23 %) were the highest between
the seventeen eukaryotic genomes analysed in the study by Zemach et al., 2010, with the second
highest belonging to rice with a CHH methylation level around 5 %.
Figure 7: Cytosine methylation distribution across Physcomitrella patens genome. Genes (A) or repeats
(B) were aligned and the average methylation levels for each 100 nt intervals are plotted. The dashed lines at zero
represent the points of alignment. A: Cytosine methylation distribution at genes, wherein a decrease in all
methylation contexts is observed. B: Cytosine methylation at P. patens repeats, with high levels of all types of
cytosine methylation (Zemach et al. 2010).
In 2015, Huang et al. described an ancient origin for Pol IV and Pol V, in a phylogenetic study
involving several green algae and plant species, ranging from Chlamydomonas reinhardtii to A. thaliana,
and suggested that a nearly complete and functional RdDM pathway could have existed in the earliest
land plants. In this study the authors were able to identify clear orthologs of DCL3 and AGO4 and two
NRPE1 sequences in P. patens (Huang et al., 2015). In another study, P. patens was shown to require
DCL3 to accumulate 22, 23 and 24 nt RNAs in order to repress TEs and allow for proper development
(Cho et al., 2008). In 2015, using small RNA-sequencing techniques, Coruh and co-workers were able
to identify several loci responsible for the origin of siRNAs ranging from 20-22 nt and from 23-24 nt
15
(heterochromatic siRNAs) and to link this origin to the 5-mC distribution across the genome of this plant.
They found that microRNAs and to a minor extend siRNAs with sizes from 20- 22 nt were derived from
regions with low 5-mC density and showed some tendency to overlap with genes, while heterochromatic
siRNAs (23-24 nt siRNAs) were associated with intergenic regions, repeats and regions with dense DNA
methylation in all sequence contexts. By mutant analysis the authors also concluded that P. patens’
heterochromatic siRNAs have a largely similar biogenesis pathway as in flowering plants, but diverge
on the relative levels of expression in vegetative tissues and on the fact that P. patens uses a novel
mDLC (minimal dicer-like) protein in conjunction with DCL3 to produce 23 nt siRNAs (Coruh et al., 2015).
To date, no evidences of deposition of 5-mC directed by siRNAs were reported for P. patens.
In a study published in 2012, Malik et al. used zebularine (a known DMTase inhibitor) to study the
effects of a genome-wide loss of 5-mC on P. patens physiology and development. In rice, this drug is
known to cause stable genomic demethylation patterns and dwarfing of plants, while in A. thaliana
seedlings, a reduced seedling growth and demethylation of repetitive sequences were observed
(Baubec et al., 2009; Zhou et al., 2002). By using P. patens, the authors detected that between 10 %
and 50 % of all 5-mC present in gametophytes was lost in plants exposed to 40 and 160 μM of
zebularine, respectively. The plants showed a reduction in the size of the gametophores that had more
elongated cells and less chloroplasts and a delayed differentiation of chloronema cells was also
observed. This phenotype could be restored upon rescuing the plants in normal medium during 30 days.
Therefore, the authors concluded that DNA methylation levels have a profound effect on growth and
differentiation of cells during gametophyte development in P. patens (Malik et al., 2012). In the same
study, the authors used an in silico approach to detect genes encoding DMTases in the P. patens
genome, which revealed the presence of seven loci possibly encoding such enzymes (Table 1): five of
those genes appear to code for methyltransferases homologous to the ones present in flowering plants,
while two others appear to be related to the human DNMT3a (Pp1s52_118V6.1) and DNMT3b
(Pp1s1_561V6.1) methyltransferases (Malik et al., 2012) (Table 1).
P. patens is the earliest diverged plant in which a CMT gene was identifed, knock-out (K.O.)
mutations of this gene causes developmental deffects, such as smaller plants, cell division rates up to
5 times slower than normal (due to the alteration of gene expression of genes involved in actin
localization), arrest of protonema growth, the absence of gametangia and, accordingly, sporophytes.
Overal, CMT mutants revealed genome-wide hypomethylation with an almost complete loss of CHG
methylation, while no significant changes in CG methylation were observed. In loci rich in CHG
methylation a partial (~ 22 %) reduction of CHH methylation was detected but this drop was not detected
for loci rich only in CHH methylation (Dangwal et al., 2014; Noy-Malka et al., 2014). More recently,
microarray analysis was performed on cmt mutants which, due to the upregulation of repetitive
sequences, allowed the authors to conclude that CMT regulates repetitive sequences that are highly
methylated (60-80 % CG, 60-80 % CHG and 30-40 % CHH) in the P. patens genome (Yaari et al., 2015).
Therefore, P. patens’ CMT is essential for normal development through gene expression regulation,
silencing of repetitive sequences regulation and maintenance of genome-wide 5-mC distribution.
16
Table 1: DNA methyltransferase (DMTases) genes present in the P. patens genome. Gene identification based on genome version 1.6. Gene name accordingly to Malik et al., 2012 and the DMTase (from A. thaliana (At) or human (Hs) with homology to the predicted protein encoded by each gene.
Gene identification in P. patens genome Gene name in P. patens Homologous DNA methyltransferase
Pp1s31_379V6.1 MET1 MET1 (At)
Pp1s117_71V6.1 CMT CMT (At)
Pp1s128_120V6.1 DNMT2 DNMT2 (At)
Pp1s271_1V6.1 DRM1 DRM1 (At)
Pp1s104_134V6.1 DRM2 DRM2 (At)
Pp1s52_118V6.1 DNMT3a DNMT3a (Hs)
Pp1s1_561V6.1 DNMT3b DNMT3b (Hs)
In 2015, Yaari and co-workers studied the function of P. patens MET1, showing a dramatic loss
of CG methylation and a reduction of CHG methylation only at CCG sites (that means on H representing
a cytosine nucleotide) when the MET1 gene was disrupted. CHH methylation levels were somewhat
decreased overall, except in loci enriched only for CHH methylation. This indicates that Physcomitrella
patens MET1 is involved in methylation of CG and CCG sites, but not at CHH rich loci. Physiologically
P. patens MET1 mutant plants develop normally but they fail to form sporophytes, indicating that MET1
is not essential for vegetative development of P. patens, but it may have an essential role in either
gamete formation, fertilization or sporophyte development. As for CMT mutants, MET1 mutant
microarray results also detected upregulation of a subset of repetitive sequences, suggesting that both
enzymes can cooperate to silence these repetitive sequences. The authors also reported that they were
not able to generate double mutants for these two genes and present their overlapping function as a
possible reason for the failure to obtain the double mutants (Yaari et al., 2015).
Physcomitrella patens transcriptomic atlas and specific expression of DNA methyltransferases
Recently, microarray analysis of the different tissues of P. patens, covering both the vegetative
stages: protonema (caulonema and chloronema), rhizoids and gametophores, as well as the
reproductive phases of development: antheridia, archegonia, antherozoids (or sperm cells), the different
stages of sporophyte development (S1 to SM, mature sporophyte) and the spores, were used to generate
a transcriptome atlas of this plant (Hernández-Coronado, 2015; Ortiz-Ramírez et al.).
This transcriptome atlas was used to explore the expression of the different DMTase genes in P.
patens’ genome (Table 2). Whereas some DMTases genes are only expressed in a few developmental
stages (e.g. MET1), others are expressed during most part of P. patens life cycle (e.g. CMT) (Table 2,
Hernández-Coronado, 2015; Ortiz-Ramírez et al.).
In what concerns the maintenance DMTases, MET1 seems to be expressed only in the
archegonia and the S2 stage of sporophyte development, while CMT is only absent in the antherozoids
and the SM phase (Table 2, Hernández-Coronado, 2015; Ortiz-Ramírez et al.). This data seems to
corroborate the phenotypes reported for CMT and MET1 deletion mutants wherein CMT mutants show
defects in vegetative growth, cell division and gametangia formation (Noy-Malka et al., 2014), being
17
expressed in all of these tissues (Table 2, Hernández-Coronado, 2015; Ortiz-Ramírez et al.). MET1
mutants develop normally but no sporophytes are formed (Yaari et al., 2015) and MET1 expression was
only detected in archegonia and S2 stage of sporophyte development (Table 2, Hernández-Coronado,
2015; Ortiz-Ramírez et al.).
Table 2: Presence and absence call for the expression of Physcomitrella patens DNA
methyltransferases genes. Gene identification from version 1.6 of the genome, the dots represent expression detected and the absence of dots means that no significant expression level was detected. Red dots represent maintenance DMTase gene MET1; Purple dots represent CMT gene; Blue dots represent de novo DMTase genes with homology to A. thaliana’s: DRM1 (dark blue) and DRM2 (lighter blue); Green dots represent de novo DMTases genes with homology to the human ones: DNMT3a (dark green) and DNMT3b (lighter green) (Adapted from Hernández-Coronado, 2015).
As for the de novo DMTases, DRM1 and DRM2 expression patterns seem to complement each
other, since DRM1 is expressed in all the tissues except the antherozoids (SCs) and DRM2 transcripts
are only detected in these gametes. DNMT3a seems to have the same expression profile as DRM1
being expressed in all tissues analyzed except the SCs, while DNMT3b transcripts are detected in the
antherozoids and the S3 stage of sporophyte development (Table 2, Hernández-Coronado, 2015; Ortiz-
Ramírez et al.)
As above mentioned, the data used for the transcriptome atlas involved samples from P. patens
antherozoids. These cells are released in clusters of approximately 50 to 150 cells and, in order to collect
enough antherozoids for transcriptomic analysis about 200-400 clusters per sample were required. In
order to obtain these samples, the method used was based on the manually dissection of mature
antheridia that were then placed on water, until the antherozoid clusters were released. The collection
of each individual cluster was achieved under an inverted microscope with a micromanipulator. This
was a time-consuming process used by Marcela Coronado to obtain the desired samples for the
transcriptomic analysis (Hernández-Coronado, 2015).
18
Aims of this study
We aim to better understand the epigenetic mechanisms acting during plant reproduction using
P. patens as our model organism. Our focus is on the antherozoid, which contribute directly to the next
generation.
Having in mind the transcriptomic profile of the DMTase genes throughout P. patens
development, it was evident that only DRM2 and DNMT3b are significantly expressed in the
antherozoids. All of these genes code for de novo methyltransferases, suggesting an important role for
de novo methylation during P. patens sexual reproduction (Table 2, Hernández-Coronado, 2015) which
may be helpful in protecting the genome of such cells from damaging events (e.g. TE insertions).
Therefore, these genes represent good candidates to help us unravel the role of de novo
methylation in P. patens, as well as to give insights into the possible reprogramming events occurring
during P. patens reproduction.
With this study we aimed to:
• Characterize the two independent K.O. mutant lines for DRM2 (Δdrm2#1 and Δdrm2#2)
previously generated in our laboratory, namely their fertilization rate and colony growth.
• Generate DNMT3b K.O. mutant lines (Δdnmt3b).
• Develop a time-efficient method to collect P. patens antherozoids by fluorescence-activated
cell sorting (FACS).
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Materials and Methods
Physcomitrella patens maintenance and growth
All Physcomitrella patens lines used in this work were maintained by vegetative propagation,
grown in solid KNOPS+T media (Table 3) for 6 to 7 days, after which they were transferred either to
new petri dishes with solid KNOPS+T media, to jiffies (small amount of highly nutritive soil, held together
by a fine netting) or stored at 4 ºC (to stop their growth). To solidify the KNOPS+T media, 7 g/L of
Formedium Agar were added to the liquid media previously prepared according to Table 3, before
autoclaving. Tissue grown on plates with solid KNOPS+T media was kept at 25 ºC with 16 hours on
light and 8 hours on dark daily cycles.
Table 3: KNOPS media constitution with the nutrients supplied to support Physcomitrella patens tissue
growth. Ammonium tartrate ((NH4)2C4H4O6) is added to compose KNOPS+T media. Glucose was added in order to compose KNOPS+GT media and Hygromycin B (Alfa Aesar) was used to constitute the KNOPS+GT+H media (grey shade).
Media component Concentration
KNOPS media
Macronutrients
CaN2O6 · 4H2O 3.39 mM
MgSO4 · 7H2O 1.01 mM
FeSO4 · 7H2O 0.04 mM
Phosphate buffer KH2PO4 (pH 6.5) 1.84 mM
Micronutrients
CuSO4 ∙ 5H2O 0.22 µM
ZnSO4 ∙ 7H2O 0.19 µM
H3BO3 9.93 µM
MnCl2 ∙ 4H2O 1.97 µM
CoCl2 ∙ 6H2O 0.23 µM
KI 0.17 µM
Na2MoO4 ∙ 2H2O 0.10 µM
KNOPS+T media Nitrogen source (NH4)2C4H4O6 2.72 mM
KNOPS+GT media Sugar source Glucose 27.75 mM
KNOPS+GT+H Selective antibiotic Hygromycin B (Alfa Aesar) 47.4 µM
The jiffies were grouped in boxes of four, two with wild-type (WT) (Gransden 2004 strain) tissue
and the other two with either Δdrm2#1 or Δdrm2#2 (previously obtained by Marcela Coronado and Anna
Thamm), in order to allow their full development and mimicking more natural conditions. Jiffies were
kept under the same conditions as the tissue plates during three weeks, after which the reproductive
stage of P. patens life cycle was induced. This was achieved by changing the conditions to 17 °C under
short-day conditions comprising 8 hours of light and 16 hours of dark cycles. The plants were maintained
in these conditions until they completed their life cycle (8 weeks after the induction of sexual
20
reproduction) (Cove, 2005; Strotbek et al., 2013). Jiffies were watered when required to keep a high
level of humidity.
KNOPS media (without any nitrogen source) was only used for spore germination and colony
growth assays while KNOPS+GT and KNOPS+GT+H (KNOPS+GT media supplemented with 25 mg/L
of Hygromycin B from Alfa Aesar) media were only used for growth and selection of DNMT3b K.O.
transformants, respectively (Table 3). See next section for details on these particular growth assays.
Confirmation of DRM2 deletion in the mutant lines by PCR
DNA extractions of protonemal tissue were performed by collecting 7-day old protonema tissue
from wild-type (WT), Δdrm2#1 and Δdrm2#2 lines (DRM2 K.O. mutant lines), from a KNOPS+T (Table
3) plate, followed by its immediate immersion in liquid nitrogen and storage at -80 ºC until required.
Extractions of DNA were performed using Epicenter MasterPure™ DNA purification kit (Epicentre),
following manufacturer’s instructions. Final DNA concentrations were quantified using Qubit ® 2.0
Fluorometer with the dsDNA BR (Broad range) assay kit, DNA purity was evaluated using Nanodrop
1000 (ThermoScientific) and DNA quality was assessed in a 1% agarose gel.
The confirmation of the DRM2 gene deletion was performed following a multiplex polymerase
chain reaction (PCR) approach, wherein two different set of reactions (A and B), that allowed the
distinction between WT and DRM2 K.O. based on the size of the band amplified, were performed. All
reactions were performed for the WT, Δdrm2#1 and Δdrm2#2 DNA samples in 20 µL of final volume,
consisting of: 2 µL of 10x DreamTaq™ Buffer (ThermoScientific), 0.4 µL of dNTPs (10 mM,
ThermoScientific), 0.4 µL of each primer (100 µM), 0.4 µL of DreamTaq™ DNA polymerase (5 U/µL)
(ThermoScientific), 2 µL of template DNA (20 ng/µL) and deionized water (to set the final volume to 20
µL). Reactions A used the primers named AT24, AT25 and AT26 and reactions B had the primers named
AT18, AT23, AT24 and AT27 (primer sequences can be found in Table S1).
Following an initial denaturation at 95 ºC for 3 min, amplification was performed in 35 cycles of:
95 ºC for 30 sec, annealing temperatures of 58 ºC for reactions A and 56 ºC for reactions B for 45 sec
and 72 ºC for 2 min and ending with a final extension of 10 min at 72 ºC. Amplified fragments were
separated by electrophoresis at 120 volts during 40 min in a 1% agarose gel stained with 1x RedSafe™
(iNtRON), using 5 µL of 1kb DNA ladder (NEB) and 15 µL per sample, for fragment size determination.
Agarose gel was imaged using GelDoc™ (BioRad).
Reactions using only one pair of primers were used to only one fragment and confirm the multiplex
results. Reactions to confirm the absence of the DRM2 gene in Δdrm2#1 and Δdrm2#2 lines used
primers AT24 and AT26 (reactions A). To confirm the correct integration of the 5’ and the 3’ flanking
regions the pair of primers used were: AT26 + AT25 and AT18 + AT27 (reactions B and C), respectively.
Primers CR7 and CR8 were used to confirm the presence of the antibiotic resistance gene (reactions
D). All reactions were performed with an annealing temperature of 58 ºC and amplified fragments were
separated in the same way as the multiplex reactions’ fragments.
21
Fertilization rate assessment
Fertilization rate was assessed using plants grown in jiffies 6 weeks after induction of sexual
reproduction. In order to obtain a statistical significant sample size, 100 gametangia from each line were
counted and the number of sporophytes present was determined. This number represents the
percentage of fertilization events in that sample. The counts were made using a stereoscope due to the
small size of the plant (~1 cm with a ~2 mm mature sporophyte diameter). A total of 8 counts were done
per mutant line (Δdrm2#1 and Δdrm2#2) plus respective WT in the generation zero (F0) and a total of 5
counts per mutant line plus the respective WT in the generation one (F1) and generation two (F2). WT
percentages below 40% were discarded since they represent abnormal samples (Marcela Coronado,
personal communication) and replaced by new samples.
Prism5 (GraphPad) software was used for the statistical analysis of all the samples. T-tests using
the Mann-Whitney post-test for all the pairs of samples were performed considering a 95 % confidence
interval in order to evaluate possible differences between the samples’ fertilization rates.
Sporophyte collection and spore sterilization
In order to obtain the first generation (F1) tissue it was required to germinate the F0 spores of the
DRM2 K.O. lines and the respective WT, which is the generation that results from transformation. In the
same way, we germinated the F1 generation spores in order to obtain F2 generation tissue. The spores
are kept inside the sporophyte, therefore the easiest way to collect them is to collect the sporophyte.
For spore sterilization three mature sporophytes were collected into a 1.5 mL eppendorf tube and
transferred to a flow hood wherein, 1 mL of 5 % bleach solution (autoclaved) was added to the
sporophytes and incubated at room temperature for 5 minutes. The bleach solution was then removed
from the tubes that were washed 3 to 4 times with 1 mL of sterile water, without bursting the sporophytes.
Then 1 mL of sterile water was added to the washed and the sterile sporophytes were broken with a 1
mL pipette tip, releasing the spores into the water. The estimated concentration of spores was of 15
spores per microliter of water, and these were germinated in petri dishes with KNOPS media (Table 3)
or stored at 4 ºC.
DRM2 K.O. and WT spores germination and colony growth assays
For the germination of the spores, 3 μL of sterilized spores-containing water were added to a
small water bottle containing 9 mL of sterile MiliQ water, mixed and then 3 mL were distributed by each
petri dish with KNOPS media (Table 3), in order for each plate to have approximately 15 spores. This
step is conducted under a flow hood, as the spores’ sterilization in order to maintain sterile conditions
and avoid sample contamination. The plates with the spores were incubated at 25 ºC with 16 hours on
light and 8 hours on dark daily cycles during 21 days.
Colony growth after cold storage of spores
In the first germination of spores after 14 weeks of storage at 4 ºC, only three plates per line (WT,
Δdrm2#1 and Δdrm2#2) were used, and the colonies grew for 21 days before the pictures were obtained
22
using a Leica Stereoscope (LED5000 RL) with a color camera. Image analysis was performed using
imageJ software (http://imagej.nih.gov/ij/). A more detailed assay was performed using new sporophyte
samples collected from WT, Δdrm2#1 and Δdrm2#2 lines and spores were sterilized as above
described, germinated on 10 KNOPS plates (Table 3) per line (fresh colony samples) and then stored
at 4 ºC.
Every two weeks and until the 14th week of cold storage, those spore samples were used to
germinate spores of each line (spores with 2, 4, 6, 8, 10, 12 and 14 weeks of cold storage). The growth
of the colonies was followed by imaging of the colonies’ autofluorescence (Texas Red (TxRed) channel)
using a Zeiss Stereo LUMAR stereoscope controlled with MicroManager version 1.14 software, at
different days of their growth: days 3, 5, 7, 10, 15 and finally day 21 of growth when, and whenever
possible, 25 colonies were picked for colony dry weight assessment. Plants from WT#1 and Δdrm2#1
were grown together, but apart from WT#2 and Δdrm2#2 plants (that were also grown together).
Sporophytes from plants grown together, were collected, sterilized and germinated at the same time
therefore, samples from WT#1 and Δdrm2#1 are comparable (as samples from WT#2 and Δdrm2#2)
and were analyzed separately from WT#2 and Δdrm2#2 lines.
Colony area measurement and analysis
All images of autofluorescence obtained from the growth of colonies were manually screened for
image quality and only good quality and clean images (without dust, debris or agar shades), were used
in further analyses. Colony area was determined using ImageJ software and the following steps (Figure
8): first, the sequences from the same sample, day of analysis and obtained using the same amplification
were imported as an image sequence (Figure 8 A) and a global scale based on the amplification of the
images was set. Next, a filter with a Gaussian blur sigma value of 6.00 (scaled units) was used for each
image (Figure 8 B). Afterwards, the threshold for each image was calculated, using Huang’s threshold
option (Figure 8 C), followed by the manual verification of the intensity of the moss (255 intensity units)
and the absence of any other signals besides the cellular autofluorescence. Finally the measurements
were conducted in ImageJ software, analyzing the particles of each image with a pixel-intensity higher
than 100 and measuring the sum of the area of those particles in each image in scaled units (total area
of the colony) (Figure 8 D).
All the data was then statistically analyzed in Prism5 (GraphPad) software. Analysis between WT
(WT#1 and WT#2) and DRM2 K.O. lines (Δdrm2#1 and Δdrm2#2) samples with equivalent time of cold
storage and day of growth were compared by t-test with Mann-Whitney post-test (not assuming normal
distribution of the samples), in order to evaluate possible differences due to the deletion of the DRM2
gene.
In order to detect possible effects of the cold storage of spores among the same line’s samples,
comparisons among the same line’s samples at the same day of growth, but different times of spores’
storage at 4 ºC was performed by one-way analysis of variance (ANOVA) using Kruskal-Wallis test and
Dunn’s multiple comparison test since some of the samples did not follow a Gaussian distribution.
23
Figure 8: Process of the measurement of the colony’s area in ImageJ software. A: Original image of the autofluorescence of a colony, obtained with Texas Red channel in a Zeiss Stereo LUMAR stereoscope controlled with MicroManager version 1.14 software. B: Image after the Gaussian blur step of the analysis. C: Image after Huang’s threshold was applied to the original image (A). D: Table with the results of the analysis of several images, wherein the colonies’ area are obtained.
Colonies dry weight and analysis
After 21 days of growth, 25 colonies per sample (per line and time of spore storage at 4 ºC) were
picked from their plates in order to access their dry weight. After removal of the colonies from the plate,
these were placed in 2 mL eppendorf tubes with 300 µL of a solution with 1.0 M NaCl, 50 mM MOPS
buffer at pH 7.0 and 15 % isopropanol (v/v) and where incubated at 65 ºC for 5 min to melt all the agar
attached to the colonies. The colonies were then dried using weighted paper pieces and a microwave
oven: colonies were placed in a previously weighted paper piece and microwaved for 2 minutes after
which the samples (colony + paper) were weighted again, this step was repeated until 2 consecutive
measurements were similar (difference lower than 0.1 mg).
The final colony weights were determined by subtracting the paper weight from the final sample
weight and the data was analyzed statistically using Prism5 (GraphPad) software. Mann-Whitney t-tests
were performed for each WT and respective mutant line pair of samples (this means for each cold
storage time of spores), considering a 95 % confidence interval, in order to detect possible differences
between the weight of the WT and the mutant colonies that originated from spores stored at 4 ºC for the
same period of time.
Kruskal-Wallis’ one-way analysis of variance (ANOVA) followed by Dunn’s multiple comparison
tests were performed between all the WT#1, WT#2, Δdrm2#1 and Δdrm2#2 samples individually, in
order to evaluate the effects of cold storage per sample type.
WT spore germination and colony growth under different pH conditions
For the germination of the WT spores and the growth of the colonies under different pH conditions,
new sporophytes were collected and spores were germinated. About 6 μL of water-containing sterilized
spores were added to a small bottle, containing 9 mL of buffered and autoclaved water at different pH
values, mixed and then divided by 5 petri dishes (3 mL to each plate) with KNOPS media (Table 3), in
order for each plate to have approximately 15 spores. All of these steps were performed under a flow
hood to keep sterile conditions. The plates with the spores were incubated at 25 ºC with 16 hours on
light and 8 hours on dark daily cycles during 21 days, when pictures were obtained using the a Leica
24
Stereoscope (LED5000 RL) with a color camera and image analysis was performed using imageJ
software (http://imagej.nih.gov/ij/).
Six different pH values were used in this assay: 6.52, 6.9, 7.51, 7.8, 8.28 and 8.93. The bottles
with buffered water at pH values of 6.52, 6.9 and 7.51 contained 5 mM MOPS buffer while the bottles
with the water at pH 7.8, 8.28 and 8.93 were buffered with 5 mM of Tris-HCl.
pSP3b plasmid construction
The plasmid used to obtain the DNMT3b K.O. mutant lines was engineered on the basis of the
plasmid pBH. This plasmid contains an ampicillin resistance marker for bacteria and hygromycin B
resistance gene for transformant selection in planta. Upstream of the hygromycin resistance gene,
1118bp of the 5’ flanking region of the P. patens DNMT3b gene were inserted, as well as 1180bp of the
3’ flaking region of this same gene, downstream of this marker in order to recombine with their genomic
homologs when inserted into the protoplasts during the transformation. After the 5’ flanking region insert,
mCherry (a sub-type of a red fluorescent protein - RFP protein) coding sequence was inserted in order
for its expression to be driven by the DNMT3b promoter (Figure 9).
First, 1.5 µg of pBH plasmid DNA (~5 µL) was digested with 1 µL XbaI restriction enzyme (5 U/µL,
NEB) and 2 µL KpnI-HF (High-Fidelity) restriction enzyme (5 U/µL, NEB) in a reaction with 3 µL of 10x
CutSmart ® Buffer (NEB) and 19 µL deionized water, making the reaction final volume of 30 µL. Plasmid
digestion proceeded at 37 ºC for 10 h, followed by enzyme’s inactivation at 65 ºC for 20 min. Afterwards,
1 µL of Antarctic Phosphatase enzyme (5 U/µL, NEB) and 3.5 µL of 10x Antarctic Phosphatase Reaction
Buffer (NEB) were added to the inactivated digestion reaction and incubated at 37 ºC for 30 min, followed
by inactivation at 70 ºC for 5 min. Digested plasmid was loaded into a 1% agarose gel stained with 1x
GelRed™ (Biotium) and the fragments were separated at 100 volts for 1 h. The gel was observed using
GelDoc™ (BioRad). The band corresponding to the digested plasmid was removed from the gel and its
DNA purified using ZymoClean™ Gel DNA Recovery kit (Zymo Research) according to manufacturer
instructions.
The 3’ end flanking region was amplified in a 50 µL PCR reactions consisting of: 10 µL of 5x
Phusion ® HF Reaction Buffer (NEB), 1 µL of dNTPs (10 mM, ThermoScientific), 2.5 µL of each primer
(100 µM, Table S1), 0.5 µL of Phusion ® High-Fidelity DNA Polymerase (5 U/µL) (NEB), 2 µL of template
DNA (20 ng/µL) and deionized water (to set the final volume to 50 µL). Amplification of the desired
fragment was achieved by an initial denaturation at 95 ºC for 3 min, followed by 35 cycles of: 95 ºC for
30 sec, 56 ºC (annealing) for 30 sec and 72 ºC for 1 min, ending with a final extension of 10 min at 72
ºC. 5 µL of the PCR products were run (100 volts for 30 min) on 1% agarose gels stained with 1x
RedSafe™ (iNtRON) using 5 µL of 1kb DNA ladder (NEB), to confirm band size. The remaining 45 µL
of the PCR products were purified using NucleoSpin ® Gel and PCR Clean-up kit (Macherey-Nagel)
following manufacturer instructions with elution in 20 µL of NE buffer.
To ensure insert’s proper orientation, specific restriction enzyme cutting sites were added to each
primer (Table S1). 15 µL of eluted fragment were digested with 1 µL XbaI restriction enzyme (5 U/µL,
NEB) and 2 µL KpnI-HF restriction enzyme (5 U/µL, NEB) in a reaction with 3 µL of 10x CutSmart ®
Buffer (NEB) and 9 µL deionized water, making the reaction final volume of 30 µL. Digestion occurred
25
at 37 ºC for 5 h, followed by enzyme’s inactivation at 65 ºC for 20 min. Digested fragment was loaded
into a 1% agarose gel stained with 1x GelRed™ (Biotium) and the sample separated at 100 volts for 1
h and observed using GelDoc™ (BioRad). The band corresponding to the digested insert was taken
from the gel and the DNA purified using ZymoClean™ Gel DNA Recovery kit (Zymo Research)
according to manufacturer instructions.
Insert’s ligation into the digested vector was achieved in a 10 µL reaction composed of: 30 ng of
plasmid (1.64 µL), 17 ng of insert (1.5 µL), 0.5 µL of T4 DNA Ligase (400 U/µL, NEB), 1 µL of 10x T4
DNA Ligase Reaction Buffer (NEB) and 5.36 µL of deionized water. This reaction was incubated
overnight at 14 ºC, followed by ligation inactivation at 65 ºC for 10 min. Transformation of Escherichia
coli (E. coli) MACH1 competent cells was achieved by adding 5 µL of the ligation reaction to 50 µL of
the competent cells, incubation for 5 min at 4 ºC followed by a heat shock at 42 ºC for 1 min. Cells were
immediately transferred to ice for 15 min, followed by the addition of 250 µL of Luria Broth (LB) media
(10 g/L tryptone, 5 g/L yeast extract, 5 g/L NaCl). Next, cells were incubated at 37 ºC with 180 rpm
shaking for 1 h after which 150, 100 and 50 µL of the cells were plated into LB plates with 100 µg/mL
ampicillin and kept at 37 ºC for 16 h.
A total of 15 colonies were selected and transferred to 10 µL of deionized water and incubated at
95 ºC for 10 min. Afterwards, 20 µL PCR reactions consisting of: 2 µL of 10x DreamTaq™ Buffer
(ThermoScientific), 0.2 µL of dNTPs (10 mM, ThermoScientific), 0.4 µL of each primer (100 µM), 0.2 µL
of DreamTaq™ DNA polymerase (5 U/µL) (ThermoScientific), 1 µL of template DNA and deionized
water (to set the final volume to 20 µL) were performed, using the primers used for the insert’s
amplification (Table S1) under the same reaction conditions. All PCR products were run on 1% agarose
gels stained with 1x RedSafe™ (iNtRON), using 5 µL of 1kb DNA ladder (NEB) at 100 volts for 30 min,
imaged with GelDoc™ (BioRad), to check for the insert’s presence. 4 colonies who appeared to have
the insert from the PCR experiment were inoculated into 5 mL of liquid LB media with 100 µg/mL
ampicillin and incubated at 37 ºC with 180 rpm shaking for 12 h.
DNA extractions from the colonies grown in liquid LB media were performed with the ZR Plasmid
Miniprep™ – Classic kit (Zymo Research), following manufacturer instructions. DNA quantification was
performed using NanoDrop 1000 (ThermoScientific). 500 ng of plasmid was digested with the insert-
specific DNA restriction enzymes (1 µL XbaI and 2 µL KpnI-HF) and with 1 µL HindIII-HF enzyme (to
linearize the plasmid, NEB) in 20 µL reactions with 2 µL of 10x CutSmart ® Buffer (NEB) and deionized
water to make the reaction final volume of 20 µL. Plasmid digestions proceeded at 37 ºC for 4 h, followed
by enzyme’s inactivation at 80 ºC for 20 min. Digested samples were separated in 1% agarose gels
stained with 1x RedSafe™ (iNtRON), using 5 µL of 1kb DNA ladder (NEB) at 100 volts for 30 min to
check for the insert’s presence and the total size of the plasmid. The agarose gel was observed using
GelDoc™ (BioRad).
All the insert sequences were confirmed by Sanger sequencing, using primers flaking the insertion
sites (Table S1). For Sanger sequencing, reactions were prepared using 2 µL of Reaction Buffer, 2 µL
of Terminator Ready Reaction Mix (obtained from the Genomics Unit at the IGC), 300 ng of total
template DNA, 0.5 µL of primer (100 mM) and deionized water so the final volume of the reaction was
10 µL. The fragment to be sequenced was amplified using the following conditions: initial denaturation
26
by rapid thermal ramp to 96 ºC and then, 96 ºC for 1 min; 25 cycles of: rapid thermal ramp to 96 ºC, 96
ºC for 10 sec, rapid thermal ramp to 50 ºC, 50 ºC for 5 sec, rapid thermal ramp to 60 ºC and 60 ºC for 4
min and finishing with a rapid thermal ramp to 4 ºC with hold until purification started. Purification was
conducted by adding to the 10 µL amplification reaction, 2 µL of a 3 M solution of sodium acetate (pH
4.6) and 50 µL of 95 % ethanol. This mixture was incubated at room temperature (RT) for 30 min followed
by centrifugation at 4 ºC at 20817 g for 30 min. Supernatant was discarded and the pellet was rinsed
with 250 µL of 70 % ethanol. Next, the sample was centrifuged at 4 ºC at 20817 g for 15 min, supernatant
was aspirated and the pellet was dried and submitted to the IGC’s Genomics Unit - sequencing service,
for sequencing. Results were aligned to the Phytozome version 10.3 corresponding sequence using the
Bioedit Sequence Alignment Editor (version 7.2.5) software.
After obtaining the plasmid with the 3’end region (pBH_3), the 5’ flanking region followed by the
mCherry coding sequence were cloned by Gibson Assembly method. This method uses primers with
sequences complementary both to the end of the region to anneal and to the end of the region where
the amplified fragment will be inserted. Therefore, 5’ flanking region forward primer was complementary
both to the region of the pBH_3 plasmid where it would be inserted and to the 5’ region to be amplified,
as its reverse primer had a sequence complementary to the end of the 5’ region to be amplified and
another region complementary to the beginning of the mCherry coding sequence where it would be
ligated. The mCherry forward primer had regions complementary to the end of the 5’ flanking region of
the DNMT3b gene and to the start of its coding sequence, and the reverse primer had sequences
annealing to the end of the mCherry coding sequence and to the end of the pBH_3 plasmid where it
would be inserted.
First 2 µg of the pBH_3 plasmid were digested with 1 µL NotI-HF (5 U/µL NEB), 1 µL PacI (5
U/µL, NEB), 2 µL of 10x CutSmart ® Buffer (NEB) and deionized water to make up the volume to a total
of 20 µL, during 9 h at 37 ºC, followed by the inactivation of the enzymes at 65 ºC for 20 min. Digested
plasmid was loaded into a 1% agarose gel stained with 1x GelRed™ (Biotium) and the separated at 100
volts for 1 h. The band corresponding to the digested plasmid was observed using GelDoc™ (BioRad),
removed from the gel and its DNA purified using ZymoClean™ Gel DNA Recovery kit (Zymo Research)
according to manufacturer instructions.
The 5’ end flanking region and the mCherry coding sequence were amplified in 50 µL PCR
reactions consisting of: 10 µL of 5x Phusion ® HF Reaction Buffer (NEB), 1 µL of dNTPs (10 mM,
ThermoScientific), 2.5 µL of each primer (100 µM, Table S1), 0.5 µL of Phusion ® High-Fidelity DNA
Polymerase (5 U/µL) (NEB), 1 µL of template DNA (20 ng/µL for 5’ region and 2 ng/µL for mCherry
sequence, where the template was another plasmid with this sequence) and deionized water (to set the
final volume to 50 µL). Amplification of the desired fragment was achieved by an initial denaturation at
95 ºC for 3 min, followed by 35 cycles of: 95 ºC for 30 sec, 56 ºC (annealing) for 30 sec and 72 ºC for 1
min, ending with a final extension of 10 min at 72 ºC. 5 µL of the PCR products were run (100 volts for
30 min) on 1 % agarose gels stained with 1x RedSafe™ (iNtRON), using 5 µL of 1kb DNA ladder (NEB),
to confirm band size. The remaining 45 µL of the PCR products were loaded into a 1 % agarose gel
stained with 1x GelRed™ (Biotium), separated at 100 volts for 1 h and imaged with GelDoc™ (BioRad).
27
The band corresponding to the digested insert was taken from the gel and the DNA purified using
ZymoClean™ Gel DNA Recovery kit (Zymo Research) following to manufacturer instructions.
Figure 9: pSP3b plasmid map (with a total of 8256 nt). The plasmid was constructed based on the pBH plasmid by the insertion the 3’ flanking region of Physcomitrella patens (P. patens) DNMT3b gene and part of the DNMT3b’ promoter (5’ flanking region) – regions in green. After DNMT3b 5’ flanking region, the mCherry coding sequence was inserted (red region) followed by the NOS terminator sequence (grey). 35S promoter (white region between red and dark blue regions) will regulate the expression of the hygromycin B resistance gene (HygR, dark blue region) and the poly(A) signal for the termination of the expression is located next (grey). This is the DNA region engineered, that will be introduced into P. patens protoplasts in order to generate the DNMT3b mutant due to the homologous recombination of the flanking regions (green). The region in yellow represents the origin of replication for bacterial cells, the white region after the yellow one is the promotor for the ampicillin resistance gene (in light blue).
Both inserts were simultaneously ligated into 40 ng of the pBH_3 digested vector. 22.6 ng of the
5’ flanking region insert and 20.4 ng of the mCherry insert were mixed with the 40 ng of the digested
vector, 2 µL of the 2x Gibson Assembly Master Mix (NEB) and deionized water was added to complete
the reaction volume to a final of 10 µL. Ligation proceeded by incubation of the sample at 50 ºC for 1 h.
28
Afterwards, 4 µL of the ligation mixture were transformed into E. coli MACH1 competent cells as
described for the cloning of the 3’ flanking region.
A total of 16 colonies were selected and transferred to 10 µL of deionized water and incubated at
95 ºC for 10 min. 20 µL PCR reactions consisting of: 2 µL of 10x DreamTaq™ Buffer (ThermoScientific),
0.2 µL of dNTPs (10 mM, ThermoScientific), 0.4 µL of each primer (100 µM), 0.2 µL of DreamTaq™
DNA polymerase (5 U/µL, ThermoScientific), 1 µL of template DNA and deionized water (to set the final
volume to 20 µL) were performed, using the forward primer from the mCherry sequence and a reverse
primer in the plasmid (Table S1). Amplification started with an initial denaturation at 95 ºC for 3 min,
followed by 35 cycles of: 95 ºC for 30 sec, 57 ºC (annealing) for 30 sec and 72 ºC for 1 min, ending with
a final extension of 10 min at 72 ºC. All of the PCR products were loaded into a 1% agarose gel stained
with 1x RedSafe™ (iNtRON) using 5 µL of 1kb DNA ladder (NEB), separated at 100 volts for 30 min, to
confirm correct fragment insertion by imaging the gel with GelDoc™ (BioRad). All colonies appeared to
have the correction insert size and only 2 were selected for liquid culture growth into 5 mL of liquid LB
media with 100 µg/mL ampicillin and incubated at 37 ºC with 180 rpm shaking for 16 h.
DNA extractions from the colonies grown in liquid LB media were performed with the ZR Plasmid
Miniprep™ – Classic kit (Zymo Research), following manufacturer instructions. DNA quantification was
performed using NanoDrop 1000 (ThermoScientific). 500 ng of plasmid was digested with the specific
DNA restriction enzymes (to allow the detection of both fragments in the plasmid): 1 µL XbaI and 1 µL
EcoRV-HF, 1 µL PacI and 1 µL EcoRV-HF and with 1 µL EcoRV-HF enzyme (all enzymes are from
NEB) in 20 µL reactions with 2 µL of 10x CutSmart ® Buffer (NEB) and deionized water to make the
reaction final volume of 20 µL. Plasmid digestions proceeded at 37 ºC for 4 h, followed by enzyme’s
inactivation at 80 ºC for 20 min. Digested samples were separated in 1 % agarose gels stained with 1x
RedSafe™ (iNtRON), using 5 µL of 1kb DNA ladder (NEB) at 100 volts for 30 min to check for the
insert’s presence and the total size of the plasmid. The gel was imaged via GelDoc™ (BioRad).
Insert’s sequences were confirmed by Sanger sequencing, using primers flaking the insertion
sites (Table S1). Sequencing samples with 500 ng of template DNA, 2.5 µL of primer (100 mM) and
deionized water to complete the reaction volume to 10 µL were submitted and the reactions were
performed by LIGHTRUN™ Sequencing Service (GATC). Sequencing results were aligned to the
Phytozome version 10.3 corresponding sequence using the Bioedit Sequence Alignment Editor (version
7.2.5) software. The final plasmid obtained was named pSP3b and its map is presented in Figure 9
(composed in SnapGene ® Version 2.8 software (GSL Biotech LLC).
Transformation of WT line with linearized pSP3b plasmid
Plant transformation was executed following a protocol adapted from (Cove, 2005; Schaefer and
Zrÿd, 2001). This protocol is divided into three main phases: first, the DNA preparation then, the plant
transformation itself and finally the protoplasts regeneration and transformants selection. The solution
compositions are detail in (Table S2).
DNA preparation: digestion and precipitation
pSP3b positive selected bacterial colony was grown in a 5 mL LB liquid media culture with 100
µg/mL ampicillin and incubated at 37 ºC with 180 rpm shaking for 12 h. Afterwards, 200 µl of the culture
29
were transferred to a 1 L Erlenmeyer containing 200 mL LB culture with 100 µg/mL ampicillin, and
incubated for 16 h at 37 ºC with 225 rpm shaking. DNA extraction of the whole culture was performed
with ZymoPure™ MaxiPrep kit, following manufacturer’s guidelines. DNA concentration was determined
using Nanodrop 1000 (ThermoScientific).
20 µl of EcoRV-HF (5 U/µl, NEB) restriction enzyme (known to cut only once in the entire
construct) was used to digest 250 µg of plasmid DNA in a reaction composed of the 20 µl of EcoRV-HF,
299 µl of the pSP3b plasmid, 500 µl of 10x CutSmart ® Buffer (NEB), 50 µl of 100x Bovine serum
albumin (BSA, NEB) and 4131 µl of deionized water. This was incubated for 15 h and 30 min at 37 ºC
followed by enzyme inactivation at 70 ºC for 20 min.
DNA precipitation was conducted by adding 400 µl of the digestion mixture (~17 µg of digested
DNA) to each tube, 13 tubes were used in total, followed by the addition of 400 µL
Phenol:Chloroform:Isoamyl Alcohol (25:24:1). The solution was mixed and centrifuged at 12000 g for
30 sec. The supernatant was collected and transferred to a new tube to which 400 µL of chloroform
were added under a fume hood. The solution was mixed and centrifuged at 12000 g for 30 sec. The
supernatant was transferred to a new tube and moved to a flow hood to keep sterile conditions.
40 µL sodium acetate (3 M, pH 5.2) and 500 µL isopropanol were added to each tube in order to
precipitate the DNA followed by a 1 h incubation at -20 ºC. The samples were centrifuged at 20817 g
for 10 min at 4 ºC. The supernatant was removed without disturbing the pellet and 1 mL of 70 % ethanol
was added to the pellet of each tube. Afterwards the samples were centrifuged for 5 min at 12000 g and
the supernatant was again discarded and the pellets were left to dry at room temperature in the flow
hood. Pellets were resuspended in 30 µl of nuclease-free water and the DNA concentration in each tube
was measured using a Nanodrop 1000 (ThermoScientific). Samples were stored at -20ºC until further
use.
Plant transformation
WT tissue from four plates after 6 days of culture was added to a petri dish containing 25 mL of 2
% Driselase ® Basidiomycetes sp. (Sigma-Aldrich) solution in 0.5 M of D-Mannitol (pH = 5.6) and
incubated for 30 min with occasional gentle mixing in a flow hood. The digested tissue sample was
filtered with a custom-made filter and funnel assembly to a beaker and left to rest for 15 min. Afterwards
the protoplasts (plant cells with digested cell wall) were slowly transferred to a 50 mL falcon tube
previously rinsed with 5 mL of a 0.5 M D-Mannitol solution and centrifuged at 88 g for 5 min. The
supernatant was carefully removed and 15 mL of the D-Mannitol solution were added in order to
resuspend the protoplasts, followed by a centrifugation (88 g, 5 min). This step was repeated, but this
time 10 mL of the D-Mannitol solution were added and a 20 µL aliquot of the protoplast sample was
used to count the protoplasts concentration using a hemocytometer under an inverted microscope
before centrifuging the sample again at 88 g for 5 min.
The protoplast sample’s supernatant was carefully removed and the volume of MMM solution
(Table S2) required to obtain 2 million protoplasts per mL was determined and added to the protoplasts’
pellet. 15 µg of precipitated DNA were added to each of the eight 50 mL falcon tubes followed by the
addition of 300 µL of the protoplast suspension in MMM solution. Then the samples were gently mixed.
30
300 µL of a 35 % polyethylene glycol 4000 (PEG) solution (Merck Milipore) (Table S2) was added, in
drops, to each falcon tube and the samples were mixed gently. A heat shock at 45 ºC for 5 min was then
applied to the samples which were then transferred to water and incubated at RT for 10 min. Afterwards,
5 mL of liquid proto solution (Table S2) were added to each tube.
The transformed protoplasts rested in the dark for 30 min at RT. Subsequently the samples were
centrifuged at 88 g for 5 min and the supernatants discarded, whereas the pellets were slowly
resuspended in 7.5 mL of liquid proto solution. To each falcon tube sample, 7.5 mL of melted top agar
(Table S2) were added and the solutions mixed. Finally, 3 mL of each sample were added to a proto-
plate (Table S2) with a cellophane membrane. All of this protocol was conducted under sterile
conditions.
Selection of stable mutant lines
The transformed protoplast proto-plates were incubated at 25 ºC under a 16 h light / 8 h dark
regime for 7 days, after which the cellophane membranes with the transformed protoplasts were
transferred to KNOPS+GT+H plates (KNOPS+GT media supplemented with 25 mg/L of hygromycin B
(Alfa Aesar, Table 3) and cultured on the same conditions for 10 days. Then the surviving colonies
(grown from regenerated protoplasts) were picked to KNOPS+GT (Table 3) plates and cultured in the
same conditions for another 10 days. Next, the surviving colonies were again transferred to
KNOPS+GT+H media plates (Table 3) and cultured for 10 days under the same conditions. Finally, a
small piece of protonema tissue from these selection-surviving colonies was collected for in-tissue PCR
reactions while the remaining tissue was plated into KNOPS+T plates (Table 3) to continue its growth,
until colony genotyping was completed.
Genotyping of potential DNMT3b K.O. lines by multiplex in-tissue PCR
For the genotyping of the lines that survived the rounds of selection after the protoplast
transformations, the confirmation of the DNMT3b gene deletion was performed by a multiplex PCR
approach directly from the tissue sample. This approach was performed using two different sets of
reactions (C and D), that allowed the distinction between the WT or Δdnmt3b colonies based on the size
of the bands amplified.
DNA samples from the selection-surviving colonies were prepared by collecting part of protonema
tissue from each colony into 30 µL of 10x DreamTaq™ Buffer (ThermoScientific) followed by immediate
freezing into liquid nitrogen. Afterwards, the samples were allowed to thaw and two more cycles of
freeze-and-thaw were performed. These samples were used as template DNA samples for the in-tissue
multiplex PCR reactions.
Reactions were performed for all of the 10 surviving colonies and a WT tissue sample as control,
in a final reaction volume of 20 µL consisting of: 5 µL of a 3 % solution of polyvinylpyrrolidone-40 (PVP-
40, Sigma-Aldrich), 0.4 µL of dNTPs (10 mM, ThermoScientific), 0.6 µL of each primer (100 µM), 0.4 µL
of DreamTaq™ DNA polymerase (5 U/µL, ThermoScientific), 2 µL of template DNA sample in 10x
DreamTaq™ Buffer (ThermoScientific) and deionized water to make up the final reaction volume.
31
Reactions C used the primers named SP31, SP32 and SP34 and reactions D had the primers
named SP30, SP31, SP33 and SP35 (primer sequences can be found on Table S1). All amplification
reactions occurred following an initial denaturation at 95 ºC for 3 min, 35 cycles of: 95 ºC for 30 sec, 55
ºC in reactions C and 58 ºC in reactions D during 45 sec and 72 ºC for 2 min and ending with a final
extension of 10 min at 72 ºC. Amplified fragments were separated by electrophoresis at 100 volts during
80 min in a 1% agarose gel stained with 1x RedSafe™ (iNtRON), using 7 µL of 1kb DNA ladder (NEB)
and 15 µL per sample, for amplified fragment size determination and genotyping assessment. All
agarose gels were imaged with GelDoc™ (BioRad).
Reactions using only one pair of primers, in order to amplify each individual fragment, were also
performed. Primers SP31 and SP32 were used in order to evaluate the absence of the DNMT3b gene
(reactions A). To confirm the correct integration of the 5’ and the 3’ flanking regions in the target genomic
region, the primers used were: SP32 + SP34 and SP33 + SP35 (reactions B and C), respectively.
Primers CR2 and CR3 were added to the reaction mix to confirm the presence of the hygromycin
resistance gene (reactions D). These reactions were all performed using the same thermocycler
program as before, but with an annealing temperature of 55 ºC. Amplified fragments were separated in
the same way as the multiplex reactions’ fragments.
Antherozoids release and labeling assays
For the development of a time-efficient method to collect P. patens sperm cells, we grew WT
tissue on jiffies for 3 weeks, under long day conditions, time after which we induced the sexual
reproduction phase. 15 days afterwards (day of sperm cell release), manual dissection of antheridia
(using a Leica Stereoscope LED3000 RL) into 20 µL of sperm-nutritive solution (Table 4, optimized by
Carlos Ramirez) was performed, during which antherozoids release occurred. The samples were then
observed under oil immersion with a 100x magnification, on a custom-built high-throughput setup, based
on a Nikon Eclipse TE2000-S equipped with a Hamamatsu Flash 2.8 sCMOS camera and controlled
with the MicroManager version 4.1.14 software.
Table 4: Sperm-nutritive solution composition. Solution optimized by Carlos Ramirez to prolong the life-time of Physcomitrella patens antherozoids and their movement.
Reagent Final concentration
CaCl2 0.450 mM
MgSO4 0.300 mM
KNO3 0.020 mM
NaHCO3 0.081 mM
The labeling of antherozoids was achieved by manually dissecting the antheridia into 20 µL of
sperm-nutritive solution pH = 6.5 (Table 4) with 10 µg/mL of fluoresceín diacetate (FDA, ThermoFisher).
Eukaryotic cell membranes are permeable to FDA that will enter the cells and will be metabolized by
cellular esterases into fluoresceín (a green fluorophore) that cannot exit the cells until they die (Jones
and Senft, 1985). The labeled samples were observed under the same microscope and with the same
32
settings as the unlabeled samples (detailed before) but also in addition, green fluorescent signal was
observed on the green fluorescent protein (GFP) channel.
Flow cytometry, cell sorting and microscopic confirmation of antherozoid samples
For the flow cytometry and cell sorting of P. patens antherozoids, mature antheridia samples were
manually dissected for 20 min into a 1.5 mL eppendorf tube with 100 µL of sperm-nutritive solution
(Table 4). Afterwards, the 100 µL sample were filtered through a custom made column + filter assembly,
wherein the filter used was a mesh filter with either 10 µm or 28 µm of pore diameter, by centrifugation
at 280 g for 1min. Next, another 100 µL of sperm nutritive solution were added to the column and filter
assembly and the centrifugation step was repeated. Finally, 100 µL of sperm nutritive solution were
added again to the column and filter assembly and centrifuged at 280 g during 1 min. This filtration steps
were designed to try to release the antherozoids from the clusters and push them through the filter, in
order to obtain them isolated in the flow-through.
A part of the flow-through sample was then analyzed in a Modular Flow Cytometer (MoFlo, Dako
Cytomation) and the cellular profile of the sample was determined (unstained control sample). Then, 0.6
µL of FDA (2 mg/µL) was added to the remaining sample and the flow cytometric analysis was repeated
(stained sample). Some of the parameters analyzed were the red signal (autofluorescence) of the cells,
measured using the FL4 (Texas Red channel) and FL7 (RFP channel) and the green signal (FDA
stained cells, viable cells), measured in the FL1 (Green) channel. The population of interest (that is
supposed to have the antherozoids) was identified by the absence of an autofluorescent (red) signal
and the presence of a FDA (green) signal. Part of this isolated population was sorted using the MoFlo
and observed using a Nikon Eclipse TE2000-S equipped with a Hamamatsu Flash 2.8 sCMOS camera
and controlled with the MicroManager version 4.1.14 software under oil immersion at 100x amplification,
in order to confirm the antherozoid presence in the analysed samples.
Phylogenetic analysis of P. patens’ de novo DNA methyltransferases genes
In order to study the evolution of Physcomitrella patens de novo methyltransferases, several
protein sequences of DNMT3a, DNMT3b, DRM1, DRM2 and DRM3 were retrieved from the NCBI
protein database (accession numbers on Table S3). P. patens protein sequences were retrieved from
Phytozome version 10.3 using the genome annotation version 3.0 and the gene identifier from Malik et
al., (2012). Identification of the 5-cytosine methyltransferase domain was performed using
InterProScan5 (EMBL-EBI) online resource and amino acid sequences of the domains were obtained
from each of the previously obtained protein sequences. Phylogenetic analyses were performed on all
sequences together (DNMT3s and DRMs), only DNMT3 and only DRM protein family sequences
separately. Analyses considering only the 5-cytosine methyltransferase domains of the sequences
obtained were also performed for all sequences (set of DNMT3s and DRMs) and for each protein family
individually (only DNMT3 and only DRM). In the analysis using the total set of sequences/domains,
sequences from Arabidopsis thaliana’s MET1, Homo sapiens’ DNMT1 and Physcomitrella patens MET1
were used as outgroups. For the analysis of DNMT3 sequences / domains P. patens DRM1 and DRM2
33
sequences were used as outgroup and when only DRM sequences / domains were considered, P.
patens DNMT3a and DNMT3b sequences were used as outgroup.
Alignments for all the set of sequences analyzed were performed using ClustalW algorithm in
Bioedit Sequence Alignment Editor (version 7.2.5) software and then were inputted as a matrix to
MEGA6 version 6.06 software. Using MEGA6, the best evolutionary model to describe the substitutions
observed in the alignments was evaluated and phylogenetic trees using the best evolutionary model per
each case, maximum likelihood methods and 500 bootstrap replications were constructed and the
consensus tree obtained.
pAT05 plasmid re-sequencing and mapping
Transformation of Escherichia coli (E. coli) MACH1 competent cells was achieved by adding 5 ng
of a sample of the plasmid used for Δdrm2 transformation (pAT05), previously obtained by Anna
Thamm, to the cells, followed by a heat shock at 42 ºC for 1 min and the transfer of the cells immediately
to ice. 250 µL of LB media were added to the sample and incubated at 37 ºC with 180 rpm shaking for
1 h after which 10, 50 and 150 µL of the cells were plated into LB plates with 100 µg/mL ampicillin and
kept at 37 ºC for 12 h. Two colonies were selected and served as inoculum for a 5 mL liquid culture of
LB media with 100 µg/mL ampicillin that grew for 12 h at 37 ºC with 180 rpm shaking. DNA extractions
from the colonies grown in liquid LB media were performed with the ZR Plasmid Miniprep™ – Classic
kit (Zymo Research), following manufacturer instructions. DNA quantification was performed using a
NanoDrop 1000 (ThermoScientific). 500 ng of plasmid was digested with several DNA restriction
enzymes for 4 h at 37 ºC to confirm the presence of the correct plasmid. Afterwards, samples were
separated in 1 % agarose gels stained with 1x RedSafe™ (iNtRON) at 120 v for 30 min to confirm the
sample’s integrity (data not shown).
DNA from one sample was also quantified using Qubit ® 2.0 Fluorometer (Invitrogen) using the
dsDNA BR assay kit (wherein BR stands for Broad Range) and a dilution to 10 ng/µL was performed
and re-measured using dsDNA HS assay kit (wherein HS stands for High Sensitivity). 20 µL of a sample
with 6.03 ng/µL were submitted to the NGS (next-generation sequencing) service at IGC. The samples
were processed according to the Illumina Nextera XT protocol, starting from the recommended 1 ng of
DNA. For library quantification it was used the quantitative PCR assay KAPA Library Quantification kit
to measure the concentration (21.4 nM) of the library and the size profile was analysed on the
Bioanalyzer (Agilent). The sample was normalized to 4 nM and pooled together with other samples,
since this only occupied 0.01 % of the Run. The pooled library was denatured and diluted, before loading
on a MiSeq cartridge (MiSeq Reagent Kit v3) for paired-end 500 cycle run (2 x 250 cycles).
After paired-end sequencing in an Illumina MiSeq, the reads obtained were sent to the
Bioinformatics unit at the IGC where assembly was performed. First, paired end reads were trimmed to
remove poor quality bases (considering a Phred Quality score < 20), using seqtk software
(https://github.com/lh3/seqtk). Filtered reads were then de novo assembled using the spades assembler
(Bankevich et al., 2012) using the set default parameters and contigs were obtained. Mapping of the
final assembled sequence was performed using SnapGene viewer ® Version 2.8 software (GSL Biotech
LLC).
34
Results
Phylogenetic analysis of Physcomitrella patens de novo methyltransferases
The phylogenetic position and conservation of de novo methyltransferases of Physcomitrella
patens were evaluated considering both the total protein sequences and only their cytosine
methyltransferase domains. Analysis of all DRM and DNMT3 sequences (total set of sequences) as well
as DRM and DNMT3 sequences separately were performed by using maximum likelihood algorithm.
The phylogenic trees obtained from the total set of sequences are shown on Figure 10. Figure 10 A
represents the tree considering the complete sequences and Figure 10 B the tree considering only the
5-mC methyltransferase domains. Trees obtained from only DRM and only DNMT3 sequences are
presented on Figure S1 and S2, respectively. All sequences accessions number are listed on Table S3
Figure 10: Phylogenetic trees obtained from the analysis of the total set of DNA methyltransferases
sequences used in this work. The cluster with DRM sequences is marked by a green rectangle, DNMT3 sequences by a blue one and DNMT1/MET1 protein sequences are highlighted by a red rectangle. Trees were obtained using maximum likelihood methods with 500 bootstrap replications and the best model to fit the alignment substitutions observed (evaluated using Mega6 software, version 6.06). The numbers in the nodes represent the percentage of trees where that branch is observed (bootstrap values), tree leafs are named by the protein name and in brackets the name of the species to whom the sequence belongs. In case of Physcomitrella patens sequences the leaves are highlighted with black dots before the leaf identifier. A: Tree obtained considering the complete protein sequences, using Jones-Taylor-Thornton model with a gamma distribution value of 4. B: Tree obtained from the sequences alignment and using the Whelan and Goldman model using a gamma distribution value of 6.
Considering the complete sequences from both DRM and DNMT3 protein families and using
Human’s DNMT1, A. thaliana’s and P. patens’ MET1 protein sequences as outgroup, the best
substitution model to fit the alignment was the Jones-Taylor-Thornton (JTT) model with a gamma
35
distribution value of 4. From the tree obtained it is possible to see a clear difference between the DNMT3,
DNMT1/MET1 and DRM sequences (Figure 10 A). P. patens’ DNMT3 sequences appear to be distinct
from the ones belonging to animals, whose DNMT3a cluster together with each other as do DNMT3b
protein sequences (Figure 10 A, blue rectangle). DNMT1/MET1 cluster with DRM sequences, being
distinct from DNMT3 proteins (Figure 10 A, red and green rectangles, respectively). All DRM sequences
cluster together, with P. patens sequences (highlighted by the black dots, Figure 10 A green rectangle)
being the most different ones and not all DRM1 cluster with each other’s, neither do DRM2 proteins as
observed for P. patens’ sequences (highlighted by the black dots, Figure 10 A).
The phylogenetic tree from the analysis of the total set of 5-mC methyltransferase domain
sequences was obtained considering the Whelan and Goldman model (WAG) using a gamma
distribution value of 6. This tree shows the sequences from DNMT1/MET1 represented as outgroup and
DNMT3, as well as DRM sequences, cluster together within its families (Figure 10 B). DNMT3 proteins
from P. patens appear to be distinct from the remaining sequences considered (highlighted by the black
dots, Figure 10 B blue rectangle), with DNMT3a from different animal species clustering together as well
as DNMT3b’s sequences. DRM1 and DRM2 sequences do not form isolated clusters and they do not
form clusters within the same species, with the exception of the Physcomitrella patens DRM1 and DRM2
sequences (highlighted by the black dots, Figure 10 B green rectangle).
Overall, both in the trees considering the total set of sequences / domains and the trees obtained
(Figure 10) from the alignments of DRM (Figure S1) and DNMT3 (Figure S2) protein sequences
separately, bootstrap values presented in the nodes are very variable and only a few are above 85-90.
P. patens DRM1 and DRM2 sequences always appear to cluster together, with bootstrap values always
above 96, and apart from the remaining DRM sequences used (Figures 10 green rectangles, S1 A and
S1 B), except when the complete DNMT3 sequences were considered and analyzed in separate (Figure
S2 A). DNMT3a and DNMT3b proteins from P. patens appear to be distinct from the remaining animal
DNMT3 sequences used for this phylogenetic inference, clustering always with each other with
bootstrap numbers always higher than 88 (Figures 10 blue rectangles, S2A and S2B) and apart from
the remaining DNMT3 sequences analyzed. Furthermore, when only DRM sequences were analyzed,
P. patens’ DNMT3 sequences formed an outgroup (Figure S1), the same occurred when DNMT3
complete sequences were analyzed (Figure S2 A) where P. patens’ DRM sequences clustered together
with the animal DNMT3 sequences while DNMT3a and DNMT3b proteins from P. patens formed a
different cluster.
Deep sequencing of pAT05 plasmid
Due to the lack of the complete sequence of the backbone plasmid that was engineered into
pAT05 plasmid by Anna Thamm and Marcela Coronado and in order to obtain the complete sequence
of the plasmid used to generate the Δdrm2 lines to be further characterized in this work, NGS
sequencing of the pAT05 plasmid was performed. From the deep sequencing of the pAT05 plasmid,
49441 reads with an average size of 863 nt, were obtained and used for the assembly of the sequence.
The de novo assembly of the reads generated a single large contig, with around 8201 nt and an
estimated coverage of about 900x. Other small contigs, with sizes lower than 300 nt and a coverage
36
between 1 and 5x, were also obtained but were considered to be artifacts from the sequencing and
assembly processes, being excluded from the mapping process. The 8201 nt contig was mapped and
the final map of the pAT05 plasmid can be seen in Figure 11.
Figure 11: Map of the complete sequence of the pAT05 (with a total of 8201 nt), used to obtain Δdrm2
lines. The plasmid was engineered by Anna Thamm and Marcela Coronado. Approximately 1000 nt of the 5’ and 3’ flanking regions of the DRM2 gene of Physcomitrella patens are represented in dark blue and where the regions designed to recombine with the genomic DNA of the wild-type in order to obtain Δdrm2 lines. After the 5’ flanking region, the eGFP (green fluorescent protein) coding sequence was inserted (green region) followed by the NOS terminator sequence (grey). 35S promoter (white region between grey and orange regions) will regulate the expression of the nptII gene (responsible by the resistance of the mutant lines to neomycin antibiotic, orange region) and the CamV poly(A) signal for the termination of the expression is located next (grey). The region in yellow represents the origin of replication for bacterial cells, the white region after the yellow one is the promotor for the ampicillin resistance gene (in light blue).
Confirmation of DRM2 deletion lines Δdrm2#1 and Δdrm2#2
In order to confirm the deletion of the DRM2 gene in Δdrm2#1 and Δdrm2#2 lines, two different
multiplex PCR reactions were performed for WT, Δdrm2#1 and Δdrm2#2 DNA samples, as well as
individual reactions of each pair of primers to amplify each individual fragment and confirm the multiplex
37
results (Figure S3). The two different reactions allowed to detect the presence/absence of the DRM2
gene (both reactions) and the presence/absence of the GFP gene (only in reactions A) and resistance
mark (only in reactions B) in the template DNA sample based on the size of the fragment amplified in
the reaction.
In Figure 12 a scheme of the strategy used for the multiplex reactions is shown and the gel
obtained by the separation of the amplified fragments is shown in Figure 13. Reactions A used primers
AT24, AT25 and AT26; the primer AT26 anneals on the genomic region near the 5’ region of the DRM2
gene, used for the homologous recombination and therefore, anneals both on WT and Δdrm2 lines
(Figure 12). Primers AT25 and AT26 allowed to distinguish between WT and K.O. samples due to the
fact that primer AT24’s complementary sequence is found in the DRM2 gene. If the transformation was
successful, DRM2 gene is only present in WT samples and reactions A should result in the amplification
of a fragment with an expected size of 2179 nt - Figure 12). In these reactions, a band around 2200 nt
is observed only in the WT sample (Figure 13, reaction A WT). AT25 primer anneals in the GFP gene
(only present in the DRM2 deletion lines) and together with primer AT24, the amplified DNA fragment is
expected to have 1278 nt (Figure 12). A band with an approximate size of 1300 nt was obtained for
samples A#1 and A#2, corresponding to reactions with Δdrm2#1 and Δdrm2#2 DNA respectively (Figure
13, reactions A#1 and A#2).
Figure 12: Scheme of the approach used for the multiplex PCR to confirm the deletion of the DRM2 gene
in Physcomitrella patens’ Δdrm2#1 and Δdrm2#2 line used in this work. On the top, a scheme of the WT genomic DNA of the DRM2 gene (yellow region) is shown, with both its promoter (5’ flanking region) and its 3’ flanking region (in blue). Below the scheme represents the same genomic DNA region as in the WT, but after the DRM2 gene deletion by homologous recombination using its 5’ and 3’ flanking regions (in blue, connected to the same regions on the WT by dashed lines). DRM2 K.O. lines have green fluorescent protein (GFP, in green) coding sequence after the 5’ flanking region of the DRM2 gene and the neomycin resistance gene (nptII R, orange), before the 3’ flanking region. Primers used: AT18 (annealing site in nptII R region), AT23 and AT24 (annealing on the DRM2 gene), AT25 (annealing in the GFP sequence), AT26 (annealing in the genomic DNA region before the recombination site), AT27 (annealing in the genomic DNA region after the recombination site), CR7 and CR8 (both specific for the nptII gene) are represented by black arrows indicating their orientation.
In reactions B the primers used were: AT18, that anneals on the neomycin phosphotransferase II
(nptII) gene, responsible for the mutant’s resistance to neomycin antibiotic (only present in the mutant
lines, Figure 12); the primer AT27, whose complementary region is on the genomic site near the 3’
38
region of the DRM2 gene (where homologous recombination took place), present on both WT and K.O.
lines (Figure 12); primer AT23 and primer AT24, both annealing at DRM2 gene-specific regions, being
only present at WT DNA (Figure 12). Primers AT18 and AT27 should only result in amplification from
the DRM2 K.O. lines’ DNA, with an expected fragment size of 2054 nt. Bands with sizes around 2000
nt were observed in the reactions B, in which Δdrm2#1 and Δdrm2#2 DNA were used as template
(Figure 13, reactions B#1 and B#2). Reactions with primers AT23 and AT24 should amplify a DNA
fragment with 935 nt from WT DNA samples (Figure 12). A band around the 900 nt region is only
detected in samples from reactions B with WT DNA (Figure 13, reaction B WT).
Figure 13: Picture of the 1 % agarose gel loaded with PCR products for wild-type (WT) and DRM2 mutant
lines (Δdrm2#1 and Δdrm2#2). The 1kb ladder (NEB) with the size of each of the ladder bands is presented on the left side of the image. A: multiplex samples from reactions A, using primers AT24, AT25 and AT26; B: multiplex samples from reactions B, using primers AT18, AT23, AT24 and AT27. WT named samples were obtained from reactions using wild-type DNA as template from amplification; #1 samples were obtained from reactions using Δdrm2#1 line’s DNA while #2 samples resulted from reactions where Δdrm2#2 line’s DNA was used as template.
In the individual reactions used to confirm the multiplex PCR results, reactions A used primers
AT24 and AT26, in order to confirm DRM2 gene’s presence (Figure 12). Samples from these reactions
show a band around the expected size (2179 nt, Figure 12) only in the reaction with WT DNA (Figure
S3, A WT). Reactions B, with AT26 and AT25 primers allowed to detect a band in the region of the
expected size one (1278 nt, Figure 12) on the Δdrm2 lines (Figure S3, B#1 and #2). In the individual
reactions C, primers AT18 and AT27 amplified a band with around 2000 nt (the expected size was of
2054 nt, Figure 12) in the Δdrm2#1 and Δdrm2#2 (samples #1 and #2) and a band with around 1000 nt
in the plasmid sample (Figure S3 C, P well). With primers CR7 and CR8 (Figure S3, reactions D), the
presence of the neomycin antibiotic was tested and a band with around 1000 nt (expected size of 1005
nt, Figure 12) was amplified in WT, Δdrm2#1 and Δdrm2#2 (WT, #1 and #2 wells respectively) samples
and a slightly bigger band in the plasmid sample (P sample). No fragments were amplified in any of the
blank samples (B well), in which water was used to replace template DNA (Figure S3).
39
Differences in fertilization rate are only detected for Δdrm2#1 in the F0 generation
Due to the previous observation that the DRM2 gene is only significantly expressed in P. patens
antherozoids, problems during sexual reproduction may occur, such as inability to fertilize the egg cell
or arrested zygote development. Therefore, the first step for the characterization of the DRM2 lines
obtained previously (Δdrm2#1 and Δdrm2#2) was to assess their fertilization rates. This assessment
was performed both for the F0 generation - tissue that was transformed as well as for the F1 and F2
generations - obtained by the germination of F0's and F1's spores respectively, for both mutant lines as
well as for the WT samples grown together with the mutant lines.
The scatter plots of the fertilization rates obtained for the each generation of the lines analysed in
this work (WT#1, WT#2, Δdrm2#1 and Δdrm2#2) are shown on Figure 14. T-tests were used in order
to assess statistically significant differences between WT#1 and Δdrm2#1 lines as well as between
WT#2 and Δdrm2#2 samples, per generation (Figure 14).
Figure 14: Fertilization rates from wild-type (WT) grown with line 1 (WT#1) and line 2 (WT#2) as well as
for DRM2 mutant lines (Δdrm2#1 and Δdrm2#2). A: Fertilization rates for generation zero (F0, n = 8); B: Fertilization rates of generation one (F1, n = 5); C: Fertilization rates of generation two (F2, n = 5). Round dots represent WT samples fertilization rate while triangles represent DRM2 mutant lines’ fertilization rates. Black horizontal bars represent the average of the lines’ fertilization rates and grey vertical lines represent samples standard error. Dashed horizontal lines represent the results of the Mann-Whitney’s t-tests performed. ns: non-significant differences; *** : p-value < 0.01, significant differences detected.
In the F0 generation (Figure 14 A), the average fertilization rate obtained for WT#1 line was of
62.88 %, this value was 62.78 % for WT#2, 47.04 % for Δdrm2#1 and for Δdrm2#2 line it was of 51.46
%. F1’s fertilization rates were 49.2 %, 53 %, 47.3 % and 50.2 % for WT#1, WT#2, Δdrm2#1 and
Δdrm2#2, respectively (Figure 14 B) and in the F2 generation average fertilization rates were 64 % for
40
WT#1, 61.2 % for WT#2, 62.6 % for Δdrm2#1 and 56.9 % for Δdrm2#2 (Figure 14 C, Table S4). Results
from the statistical analysis by t-test are summarized in Figure 14. Comparing the WT with the respective
DRM2 K.O. line, the only statistically significant difference of fertilization rate detected was between
WT#1 and Δdrm2#1 lines in the F0 generation (p-value = 0.0006, Figure 14 A). All other pairs of samples
compared showed a p-value above 0.05 and therefore, are not significantly different (Figures 14 B and
C). Values for standard deviations and standard error of all samples as well as the p-values obtained
from the statistical analysis are presented in Table S4).
Colonies appearance shows phenotypic variations after cold storage of spores
Δdrm2#2 colonies appear smaller and with a more round shape than WT colonies when
germinated from spores stored at 4 ºC during 14 weeks.
Figure 15: F0’s wild-type colonies obtained 21 days after germination of spores. Some have irregular borders (A, B, D, E and F) while others are more round shaped, having smooth borders (C), but all have started to develop gametophores (1.25x magnification). Scale bars = 1 mm.
Figure 16: F0’s Δdrm2#2 colonies obtained 21 days after germination of spores. Some have irregular borders (A - E) while others have a more regular round shape (F). All of them have already started to grow gametophores (1.25x magnification). Scale bars = 1 mm.
In order to obtain the subsequent generations (F1 and F2), the germination of freshly sterilized
F0’s spores for the WT and Δdrm2 lines analysed in this work was required. In all three lines freshly
sterilized spores germinated (WT, Δdrm2#1 and Δdrm2#2) and the colonies obtained grew
41
approximately to the same size, shape and appearance. Figures 15 and 16 show examples of 6 colonies
after 21 days of growth from WT and Δdrm2#2 line, respectively. In all samples observed, colonies with
irregular borders were detected (A, B, D, E and F in Figure 15 and A to E in Figure 16) as well as
colonies with more round shape (Figure 15 C and Figure16 F), but all of them had already started to
develop gametophores (data not shown for line Δdrm2#1).
After the storage of the same spore samples at 4 ºC for 14 weeks (3.5 months), the germination
was again repeated for the 3 lines (WT, Δdrm2#1 and Δdrm2#2). Figures 17 and 18 present examples
for colonies of F0 WT (Figure 17) and colonies of F0 Δdrm2#2 colonies obtained (Figure 18) after 21
days of growth. It is possible to see that all the WT colonies have started to develop gametophores
having irregular borders (Figure 17), while colonies from Δdrm2#2 line show a more regular round shape
and no development of gametophores (Figure 18). Furthermore, from the Δdrm2#1 line no spores
germinated (data not shown).
Figure 17: F0 wild-type colonies obtained 21 days after germination of spores kept at 4 ºC for 3.5 months. All of them have irregular borders and gametophores developing (1.25x magnification). Scale bars = 1 mm.
Figure 18: Colonies from F0 Δdrm2#2 obtained 21 days after germination of spores, kept sterilized at 4
ºC for 3.5 months. They all have a regular, round shape and none have started to develop gametophores (1.25x magnification). Scale bars = 1 mm.
42
Smaller colonies can be detected both in WT and Δdrm2 lines and do not seem to correlate with
the time of cold storage of spores
In order to further investigate the hypothesis that alterations on the methylation status of the
Δdrm2 lines lead to the observed phenotype of smaller and round shaped colonies without
gametophores, described above in the previous sub-section, a more timed and detailed assay was
required. This was achieved by analyzing the growth of colonies in dependence of the time that spores
were stored sterilized at 4 ºC.
Spores from WT#1, WT#2, Δdrm2#1 and Δdrm2#2 lines were germinated after sterilization
(freshly sterilized spores), kept at 4 ºC and then germinated again every two weeks, from 2 to 14 weeks
of cold storage.
Despite not being consistently observed again during or at the end of the colony germination
assay, smaller colonies with a round shape and no gametophores (similar to the ones observed from F0
Δdrm2#2 spores, kept sterilized at 4 ºC for 3.5 months - Figure 18) were observed on 3 out of 10 plates
from the WT#1 line after 6 weeks of cold storage of spores (Figure 19 C). For this same sample, normal
irregular colonies with gametophores were also obtained (Figure 19 A and B) in the remaining 7 of 10
plates. Colonies with different shapes were also observed only in the Δdrm2#1 line, after the spores
were stored for 10 weeks at 4 ºC. Some were irregular shaped with gametophores developing (normal
colonies) were observed in 1 plate (Figure 19 D). In another plate smaller and rounder colonies with a
few gametophores developing (Figure 19 E) were obtained. 2 out of 4 colony plates had smaller colonies
with a round shape and without gametophores (Figure 19 F). The 6 remaining plates were trashed
before the 21 days of growth were completed, due to heavy contaminations.
Figure 19: Colonies of F0 wild-type (WT), obtained from the germination of spores stored at 4 ºC during
6 weeks (A - C) and of F0 Δdrm2#1 line colonies germinated from pores stores at 4 ºC for 10 weeks, after 21
days of growth. A and B – WT colonies with a normal aspect (irregular shape and with gametophores developing); C – Small WT colony, with a round shape and without gametophores. D – Normal Δdrm2#1 colony; E – Colony with one gametophore developing from Δdrm2#1 line; F – Small Δdrm2#1 colony with no gametophores developing. All pictures were obtained using 1.25x magnification. Scale bars = 1 mm.
43
Colony growth phenotypes are not affected significantly by pH
Due to the apparently stochastic appearance of the “round shaped with no gametophores
colonies” phenotype during colony growth after cold storage of sterilized spores at 4 ºC and to variations
of pH observed in the water in the bottles used for plating the spores to germinate (varying between 6.9
and 8.9, n = 10, data not shown), one hypothesis was that this differences in pH in the water could
change the media’s pH and therefore affect the colony growth and shape. To test this, freshly sterilized
WT spores were germinated using water buffered for different pH values (pH 6.52, pH 6.90, pH 7.51,
pH 7.80 and pH 8.93).
For all pH values tested, spores germinated and 21 days-old WT colonies appeared healthy, most
of them with gametophores developing, as shown by the colonies shown on Figure 20.
Figure 20: Colonies from F0 wild-type germinated with water at different pH values, after 21 days of
growth. A: colonies germinated using water at pH 6.52; B: pictures of colonies germinated with water at pH 6.9; C: colonies germinated with water at pH 7.51; D: example of colonies germinated with water at pH 7.8; E: colonies obtained from spores germinated with water at pH 8.93. Scale bars represent 1 mm.
Growth of P. patens colonies can be followed in detail by determination of colony area and dry weight
Determination of WT and Δdrm2 colonies area and its variation with the time of cold storage of
sterilized spores.
The area of Physcomitrella patens colonies, germinated from WT#1, WT#2, Δdrm2#1 and
Δdrm2#2 spores, was evaluated during their germination and growth. Colonies were imaged at 3, 5, 7,
10, 15 and 21 days after spores being plated in order to germinate. Freshly sterilized spores as well as
spores stored at 4 ºC for 2, 4, 6, 8, 10, 12 and 14 weeks from all the lines analyzed were germinated.
44
Colony area was determined (in mm2) and the plots showing the variation of the average area form each
analyzed point can be seen in Figures 21 (lines WT#1 and Δdrm2#1) and 22 (lines WT#2 and Δdrm2#2).
Scatter plots of the area measurements in each sample can be observed in supplemental material
(Figures S4 and S5 corresponding to WT#1 samples; Figures S6 and S7 are from Δdrm2#1; Figures S8
and S9 represent the data obtained from colonies of the WT#2 line and Figures S10 and S11 area from
the colonies of Δdrm2#2 line).
Colonies area at the 3rd day of growth were not quantified, due to small size of the colonies and
the difficulty in subtracting the background noise from the samples of: WT#1 colonies from freshly
sterilized spores, spores stored at 4 ºC for 2, 4, 6 and 8 weeks and colonies of Δdrm2#1 from freshly
sterilized spores, spores stored at 4 ºC during 2 and 14 weeks. The same problem occurred for the 5th
day of growth of WT#1 colonies germinated from spores stored at 4 ºC for 2 weeks. WT#1 spores stored
during 14 weeks didn’t germinate therefore no colonies were obtained. At the 10th day of growth of
Δdrm2#1 colonies from spores stored at 4 ºC for 14 weeks the measurements failed to be obtained due
to time constraints. In all the cases where the colony area was impossible to determine, the value
attributed to those samples was zero and they were not used for statistical comparisons (Figure 21).
In Figure 21 an increase in the average colony area with the time of growth can be seen for all
samples (except for the samples with a value of zero) both for the WT#1 and Δdrm2#1 line. After 21
days of growth, the highest value of average area (35.87 mm2) obtained is from the WT#1 sample where
the spores were stored at 4 ºC for 10 weeks (WT#1 10w 4C, Figure 21, Table S5) and the lowest is from
the Δdrm2#1 14w 4C (colonies of Δdrm2#1 from the germination of spores at 4 ºC for 14 weeks, Figure
21, Table S5), with an average of 17.79 mm2. The samples from the freshly sterilized spores have almost
the lowest values of the 21 days-old colony samples with average values of 20.87 mm2 for WT#1 and
22.79 mm2 in Δdrm2#1 line (Figure 21, Table S5). The mean, standard deviation and standard error
values for all samples can be found in Table S5.
In order to assess possible differences between the growth of colonies from WT#1 and Δdrm2#1
lines at different time points, t-tests between samples of both lines obtained from spores stored in cold
for the same amount of time and at the same day of growth were executed (Table S5). Statistically
significant differences were detected between colonies from freshly sterilized spores at day 7 (p-value
= 0.0121) and day 10 of growth (p-value = 0.0126); from spores stored at 4 ºC for 2 weeks after 7 and
15 days of growth (p-values = 0.0480 and 0.0353, respectively); from spores stored for 4 weeks at 4 ºC
at day 7 and 10 of growth (p-values = 0.0155 and 0.0052, respectively). In the colonies from spores
stored at 4 ºC for 10 weeks, differences were detected after 7 (p-value = 0.0001), 10 (p-value = 0.0015),
15 (p-value = 0.0008) and 21 (p-value = 0.0004) days of growth and from spores stored during 12 weeks,
differences were detected between colonies at 5 (p-value < 0.0001), 10 (p-value = 0.0003), 7, 15 and
21 days of growth (this last three time points all had a p-value lower than 0.0001) (Table S5). Higher
values for the average colony area were detected in the Δdrm2#1 line for the freshly sterilized spores,
2 and 12 week stores spores’ samples, when compared to the WT#1 average values (Table S5).
ANOVA statistical analysis among the same line and between colony area values at the same
day of growth but whose spores were stored at 4 ºC for different periods of time were performed, in
order to evaluate possible effects of cold storage of spores in the growth of the colonies at a specific
45
growth time. Several differences were detected although none at day 3 of growth (considering WT#1 10
weeks and 12 weeks samples and Δdrm2#1 4, 6, 8, 10 and 12 weeks samples, Tables S6 and S7,
respectively).
Figure 21: Variation of average colony area with different days of growth from WT#1 and Δdrm2#1
colonies, germinated from spores stored (0 to 14 weeks) at 4 ºC. Triangles represent the average colony area of WT#1 samples and circles the average colony area from Δdrm2#1 colonies. The average values from the same sample (same time of cold storage of spores) are connected by continuous lines in WT#1 samples and dashed lines in Δdrm2#1 samples. Samples from spores stored at 4 ºC for different periods of time are represented by different colours (from dark green – freshly sterilized spores, to dark blue – from spores stored for 14 weeks). Black vertical lines represent standard error.
After 21 days of growth, the WT#1 colony areas showed differences between the samples from:
freshly sterilized spores and 2, 4 and 10 week’s stored spores, with the freshly sterilized spores colonies
Colony area of WT#1 and ∆drm2#1 colonies
Days of growth
Avera
ge c
olo
ny a
rea (
mm
2)
day 3
day 5
day 7
day 1
0
day 1
5
day 2
1
0
10
20
30
40WT#1 Fresh
WT#1 2w 4C
WT#1 4w 4C
WT#1 6w 4C
WT#1 8w 4C
WT#1 10w 4C
WT#1 12w 4C
WT#1 14w 4C
∆drm2#1 Fresh
∆drm2#1 2w 4C
∆drm2#1 4w 4C
∆drm2#1 6w 4C
∆drm2#1 8w 4C
∆drm2#1 10w 4C
∆drm2#1 12w 4C
∆drm2#1 14w 4C
46
having a lower average area value. Colonies from 2 weeks at 4 ºC spores had a significantly higher
average area value than colonies from spores stored for 6 and 12 weeks; as colonies of 4 week-stored
spores had a higher value than colonies from 6 and 12 week-stored spores. Spores stored at 4 ºC for 8
weeks gave rise to colonies with an average area higher than that of colonies from spores stored for 12
weeks (difference statistically significant). Finally, colonies from spores stored at 4 ºC for 10 weeks at
statistically significant higher average colony area than those of spores stored at 4 ºC for 6 and 12 weeks
(Tables S5 and S6)
After the same period of growth (21 days), colonies of Δdrm2#1 were considered to have different
areas between the samples from the spores stored at 4 ºC for 14 weeks and the ones from spores
stored during 2, 6 and 12 weeks (Table S7). The average colony area was always lower for the colonies
from the spores stored in cold for 14 weeks (Table S5).
The area of 3 days old colonies from WT#2 spores stored on cold for 2, 4, 6 and 8 weeks,
Δdrm2#2 freshly sterilized spores and spores stored at 4 ºC during 2, 4 and 6 weeks was not determined
due to the small size of the colonies and the high background noise of these images. Area from 15 and
21 days old colonies of WT#2 spores stored on cold for 4 weeks were not determined due to the heavy
contamination of these plates that lead them to be trashed. No colonies were obtained from spores of
WT#2 stored during 10 and 12 weeks samples (Figure 22).
Figure 22 shows an increase on the average colony area of WT#2 and Δdrm2#2 colonies, with
their growth in all samples (the values of zero were attributed when it was not possible to determine the
area values of the colonies). At 21 days of growth, the highest average area value obtained was from
the WT#2 colonies germinated from spores stored at 4 ºC for 14 weeks (31.90 mm2, Table S8) followed
by the colonies of Δdrm2#1 line obtained from 14 week-stored spores (23.64 mm2, Table S8), while the
lowest value determined was from WT#2 sample where the spores were stored at 4 ºC for 2 weeks
(WT#2 2w 4C, Figure 22, Table S8) with an average of 7.9 mm2. Colonies average area from freshly
sterilized spores after 21 days of growth for the WT#2 line (average area value of 13.98 mm2, Figure
22, Table S8) are between the lowest values detected. The opposite, with a high average colony area,
is observed for Δdrm2#2 line (21.84 mm2, Figure 22, Table S8). The mean, standard deviation values
and standard error values for all samples can be found in Table S8.
T-tests between WT#2 and Δdrm2#2 lines performed between colonies germinated from spores
stored in cold for the same time and at the same day of growth, unravel statistically significant differences
between samples from freshly sterilized spores after 7 days of growth and until 21 days of growth, with
p-values always lower than 0.0001. The same occurred for samples obtained after the spores being
stored at 4 ºC for 2 weeks, being detected after 5 days of growth (and until 21 days of growth) always
with a p-value inferior to 0.0001. The same p-values were obtained in the comparisons between the
colonies from the spores stored during 4 weeks at 4 ºC (day 5, 7 and 10) and stored for 6 weeks in the
15th day of growth. At 21 days of growth and after spores being stored for 6 weeks at 4 ºC the p-value
obtained was of 0.0105. Colonies germinated from spores stored during 8 weeks at 4 ºC were
considered different on the days 5, 7 and 15 (p-values = 0.005, 0.0002 and 0.026, respectively). After
14 weeks of cold storage of spores, the colonies obtained for the WT#2 and Δdrm2#2 lines were
47
significant different after 5 days of growth (p-values lower than 0.0001 for day 5, 7 and 10 of growth),
after 21 days of growth the p-value obtained was 0.0006 (Table S8).
Figure 22: Variation of the average colony area with the different days of growth from WT#2 and Δdrm2#2
colonies, germinated from spores stored (0 to 14 weeks) at 4 ºC. Triangles represent the average colony area of WT#2 samples and circles the average colony area from Δdrm2#2 colonies. The average values from the same sample (same time of cold storage of spores) are connected by continuous lines in WT#2 samples and dashed lines in Δdrm2#2 samples. Samples from spores stored at 4 ºC for different periods of time are represented by different colours (from dark green – freshly sterilized spores, to dark blue – from spores stored for 14 weeks). Black vertical lines represent standard error.
The area of the Δdrm2#2 colonies germinated from freshly sterilized spores, spores stored at 4ºC
for 2 and 4 weeks had always a higher average colony area higher than the WT#2 for the time points
where the differences were significant. The differences observed between the colonies from spores
Colony area of WT#2 and ∆drm2#2 colonies
Days of growth
Avera
ge c
olo
ny a
rea (
mm
2)
day 3
day 5
day 7
day 1
0
day 1
5
day 2
1
0
10
20
30
40
WT#2 Fresh
∆drm2#2 Fresh
WT#2 2w 4C
∆drm2#2 2w 4C
WT#2 4w 4C
∆drm2#2 4w 4C
WT#2 6w 4C
∆drm2#2 6w 4C
WT#2 8w 4C
∆drm2#2 8w 4C
WT#2 10w 4C
∆drm2#2 10w 4C
WT#2 12w 4C
∆drm2#2 12w 4C
WT#2 14w 4C
∆drm2#2 14w 4C
48
stored for 6 weeks showed higher average values in the WT#2 line. Comparisons among the colonies
obtained from spores stored in cold for 8 weeks showed higher average values on the Δdrm2#2 line for
the 5th and 7th days of growth than the WT#2 value, but a lower value in the 15th day of growth. In the
5th, 7th and 10th days of growth of the colonies germinated from spores stored at 4 ºC for 14 weeks higher
average colony area values were observed for the Δdrm2 line opposed to the 21st day of growth, in
which the higher value was from the WT#2 colonies (Table S8).
Among the same line samples, ANOVA statistical analysis were performed considering colony
area values at the same day of growth but whose spores were stored at 4 ºC for different periods of
time, in order to evaluate possible effects of cold storage of spores in the growth of the colonies at a
specific time-point. Several differences were detected although none at day 3 of growth for the Δdrm2#2
line (analysis between 8, 10, 12 and 14 week-stored spores, Table S10) except between freshly
sterilized spores and 8 week-stored spores for WT#2 samples (considering spores freshly sterilized and
stored for 8 and 14 weeks), but in those samples only 2 colonies were measured and the higher average
value was from the 8 week sample (Tables S8 and S9).
At 21 days of growth, differences were detected between the WT#2 colonies obtained from freshly
sterilized spores and those from spores stored at 4 ºC during 2 (lower average value than the one from
colonies from freshly sterilized spores) and 14 weeks (higher average area than the colonies from freshly
sterilized spores) and also between the colonies from spores stored in cold for 2 weeks and those from
spores stored during 6, 8 and 14 weeks, with the colonies from the 2 weeks stored spores showing a
lower average colony area than the remaining (Tables S8 and S9).
Colonies with 21 days of growth from Δdrm2#2 line also showed statistical significant differences
between colonies from freshly sterilized spores, with higher average area values, and those from spores
stored for 4, 6 and 12 weeks. Colonies from spores stored on cold for 2 weeks had a significant smaller
area than those from spores stored during 14 weeks, the same occurred for the 4 weeks stored spores’
colonies (smaller) and those germinated from spores stored for 10 and 14 weeks on cold. Colonies
grown from spores stored on cold for 6 weeks had a statistically smaller area than those from spores
stored in the same conditions during 8, 10 and 14 weeks. Samples from spores stored on cold for 12
weeks were significantly different from those arising from spores stored in the same conditions for 10
and 14 weeks, showing a consistently lower average colony area (Table S8 and S10).
All data and statistical analyses results obtained (t-tests and ANOVA) are detailed on Table S5
(WT#1 and Δdrm2#1 data and t-tests), Tables S6 and S7 (WT#1 and Δdrm2#1 ANOVA results,
respectively), Table S8 (WT#2 and Δdrm2#2 data and t-tests), Tables S9 and S10 (WT#2 and Δdrm2#2
ANOVA results, respectively).
Cold storage of spores has little effect on dry weight of WT and Δdrm2 colonies.
Another approach to assess possible differences between the germination of spores and growth
of colonies between WT and DRM2 deletion lines was to determine the dry weight of colonies from the
samples used for the colony area measurements. Whenever possible 25 colonies were analysed,
exception being the samples were no spores germinated (WT#1 14 weeks of storage, WT#2 10 and 12
weeks of 4ºC storage), which were not considered for statistical analysis either, and samples were 25
49
colonies were not obtained (Δdrm2#1 10 weeks – 17 colonies; WT#1 10 weeks – 22 colonies; Δdrm2#1
14 weeks – 13 colonies; WT#2 4 weeks – 10 colonies; WT#2 6 weeks – 20 colonies and Δdrm2#2 12
weeks of cold storage – 24 colonies).
The scatter plot with data from the WT#1 and Δdrm2#1 lines is shown in Figure 23. Distribution
of the colony weight can be seen, as well as the results from the t-tests between the WT and mutant
pairs of samples. Average colony weight, standard deviation and standard error for all the samples can
be found in Table S11, except for WT#1 colonies from spores stored at 4ºC during 14 weeks for which
no colonies were obtained and the weight attributed to this sample was set to zero. From the statistical
tests performed (t-tests – Table S11, WT#1 ANOVA – Table S13 and Δdrm2#1 ANOVA – Table S14)
no significant differences were detected.
Figure 23: Scatter plot of the dry weight of the colonies of WT#1 and Δdrm2#1 lines after 21 days of
growth, obtained from the germination of spores stored at 4 ºC for different periods of time (0 to 14 weeks). Horizontal black bars represent the colonies average dry weight (mg) and vertical grey bars represent standard error bars. Dry weight of WT#1 colonies from spores stored for 14 weeks at 4 ºC, no colonies were obtained and so the values are considered zero and were not used for statistical analysis; Circles represent colonies from WT#1 samples while triangles represent the dry weigh from the colonies of Δdrm2#1. Brackets under sample identifiers represent the comparisons evaluated by t-test analysis and their statistical result – ns means no significant differences were detected between those samples.
The data distribution of the dry weight of the colonies of WT#2 and Δdrm2#2 lines can be seen in
the scatter plot represented in Figure 24. Average dry weight, standard deviation and standard error of
all the samples are detailed in Table S12. The same statistical analysis was performed for colonies from
these lines and the results from the t-tests between the WT#2 and Δdrm2#2 pairs of samples are showed
on Figure 24. As opposed to all other samples, no colonies were obtained from the WT#2 samples
germinated after 10 and 12 weeks of spore storage. From the Mann-Whitney’s t-tests, it was only
Dry weight of colonies WT#1 and ∆drm2#1
Co
lon
y d
ry w
eig
ht
(mg
)
WT#1
Fre
sh
drm2#
1 Fre
sh
∆
WT#
1 2w
4C
drm2#
1 2w
4C
∆
WT#
1 4w
4C
drm2#
1 4w
4C
∆
WT#1
6w 4
C
drm2#
1 6w
4C
∆
WT#1
8w 4
C
drm2#
1 8w
4C
∆W
T#1 1
0w 4
C
drm2#
1 10
w 4
C
∆
WT#1
12w
4C
drm2#
1 12
w 4
C
∆
WT#1
14w
4C
drm2#
1 14
w 4
C
∆
0
5
10
15
WT#1 Fresh
∆drm 2_1 Fresh
WT#1 2w 4C
∆drm 2_1 2w 4C
WT#1 4w 4C
∆drm 2_1 4w 4C
WT#1 6w 4C
∆drm 2_1 6w 4C
WT#1 8w 4C
∆drm 2_1 8w 4C
WT#1 10w 4C
∆drm 2_1 10w 4C
WT#1 12w 4C
∆drm 2_1 12w 4C
WT#1 14w 4C
∆drm 2_1 14w 4C
50
possible to detect statistically significant differences (p-value < 0.05) between the pairs germinated after
8 (p-value = 0.0012) and 14 weeks (p-value lower than 0.0001) of cold storage of spores, with the
Δdrm2#2 values always being higher than the WT#2’s ones (Figure 24, Table S12).
Figure 24: Scatter plot of the dry weight of the WT#2 and Δdrm2#2 colonies after 21 days of growth,
obtained from the germination of sterilized spores stored at 4 ºC for different periods of time (0 to 14 weeks). Horizontal black bars represent the colonies average dry weight (mg) and vertical grey bars represent standard error bars. WT#2 10w 4C and WT#2 12w 4C – dry weight of colonies originated from spores stored at 4 ºC for 10 and 12 weeks, since no colonies were obtained, values are considered zero and were not used for statistical analysis of data. Circles represent colonies from WT#2 samples while triangles represent Δdrm2#2 colony dry weight. Brackets underneath samples’ identifiers represent the comparisons evaluated by t-test analysis and their statistical result – ns: p-value >0.05; ***: p-value <0.01; ****: p-value <0.001.
ANOVA analysis performed for the WT#2 samples showed significant differences between
colonies from spores stored at 4 ºC during 2 weeks and from spores stored for 6 and 8 weeks (with
higher 2 weeks’ values). Colonies grown from WT#2 spores stored from 4 weeks at 4 ºC were
statistically different from the colonies from spores stored for 6, 8 and 14 weeks (higher values being
observed for the 4 weeks samples) (Table S15). From the statistical analysis performed for the Δdrm2#2
lines by ANOVA, statistically significant differences were detected between colonies belonging to the
sample Δdrm2#2 4 weeks at 4 ºC and the colonies from the samples of spores stored for 6, 8 and 12
weeks, with the 4 week stored spores’ colonies showing consistently higher values (Table S16).
Δdnmt3b knockout lines were not obtained
According to the P. patens transcriptome Atlas (Hernández-Coronado, 2015; Ortiz-Ramírez et
al.), the DNMT3b gene appears to be only significantly expressed in the antherozoids and in the S3
Dry weight of colonies WT#2 and ∆drm2#2
Co
lon
y d
ry w
eig
ht
(mg
)
WT#2
Fre
sh
drm2#
2 Fre
sh
∆
WT#2
2w 4
C
drm2#
2 2w
4C
∆
WT#2
4w 4
C
drm2#
2 4w
4C
∆
WT#2
6w 4
C
drm2#
2 6w
4C
∆
WT#
2 8w
4C
drm2#
2 8w
4C
∆W
T#2 1
0w 4
C
drm2#
2 10
w 4
C
∆
WT#2
12w
4C
drm2#
2 12
w 4
C
∆
WT#2
14w
4C
drm2#
2 14
w 4
C
∆
0
5
10
15
20
WT#2 Fresh
∆drm 2_2 Fresh
WT#2 2w 4C
∆drm 2_2 2w 4C
WT#2 4w 4C
∆drm 2_2 4w 4C
WT#2 6w 4C
∆drm 2_2 6w 4C
WT#2 8w 4C
∆drm 2_2 8w 4C
WT#2 10w 4C
∆drm 2_2 10w 4C
WT#2 12w 4C
∆drm 2_2 12w 4C
WT#2 14w 4C
∆drm 2_2 14w 4C
51
stage of sporophyte development (Table 2). Both of these stages are directly related with the sexual
reproduction of P. patens and the only other de novo methyltransferase gene expressed in the
antherozoids is DRM2. From the results previously described in this work, Δdrm2 lines do not show any
strong phenotype. Possibly, DNMT3b de novo methyltransferase may compensate for the lack of DRM2
in these lines. Therefore we decided to obtain DNMT3b deletion lines.
Figure 25: Colonies obtained after two rounds of selection, regenerated from protoplasts subjected to
the transformation protocol with the pSP3b plasmid. A: Example of a selection-surviving colony. B: Example of a dead colony obtained after selection. Scale bar = 1 mm.
Figure 26: Scheme of the multiplex PCR approach used to genotype the selection-surviving colonies of
Physcomitrella patens transformed with the pSP3b plasmid. This strategy was designed to distinguish between wild-type (WT) strain colonies (top) and the Δdnmt3b lines – lower section - where the homologous recombination occurred and the DNMT3b gene was deleted. The DNMT3b gene (grey region) is only present in WT colonies, both its promoter (5’ flanking region) and its 3’ flanking region (in green) are present in WT and DNMT3b K.O. colonies since this regions were used for the homologous recombination (represented by the dashed lines). Δdnmt3b lines have the coding sequence of the mCherry fluorescent protein (in red) after the 5’ flanking region of the gene and the hygromycin B resistance gene (Hyg R, in blue), before the 3’ flanking region. Primers: SP30 and SP31 (annealing on the DNMT3b gene), SP32 (annealing in the genomic DNA region before the recombination site), SP33 (annealing in the genomic DNA region after the recombination site), SP34 (annealing in the mCherry sequence), SP35 (annealing site in Hyg R region), CR2 and CR3 (both specific for the hygromycin B resistance gene) are represented by black arrows indicating their orientation.
52
In Figure 9, the map of the plasmid pSP3b used in the transformation process can be seen. After
two rounds of selection, the colonies regenerated from the protoplasts subjected to the transformation
protocol were genotyped. A multiplex in tissue PCR strategy was used to assess the presence or
absence of the DNMT3b gene in the selection surviving colonies, allowing to distinguish WT from
Δdnmt3b genotypes by the size of the DNA fragment amplified (Figure 26). The gel obtained by the
separation of the fragments amplified from the in-tissue multiplex reactions C and D can be seen in
Figure 27. Individual reactions to amplify each individual fragment were also performed (Figure S12).
After the second round of selection on hygromycin B containing media (Table 3), only 10 surviving
colonies were obtained and 4 of them died (Figure 25), turning white due to the action of the antibiotic.
The 10 colonies (constituted by green protonema tissue) also showed some white and/or brown regions
(Figure 25). Tissue from these 10 colonies was collected into PCR reaction buffer and used for in-tissue
multiplex PCRs as well as individual PCR reactions, in order to evaluate the possible deletion of the
DNMT3b gene resulting from the homologous recombination of the plasmid with WT DNA, during the
transformation process.
Reactions C used the primers SP31 (specific for DNMT3b gene), SP32 (annealing on the genomic
DNA near the HR site, present in all samples) and SP34 (annealing in the mCherry gene present at the
pSP3b plasmid) (Figure 26). The expected sizes for the amplified fragments in the reactions C were of
2137 nt, if the DNMT3b gene is present on the sample used as template DNA (WT samples) and 1292
nt in the cases where the DNMT3b gene was efficiently replaced by the mCherry coding sequence
(Δdnmt3b lines, Figure 26).
Reactions D included 4 primers: SP30 and SP31 (DNMT3b gene specific), primer SP33
(annealing on the genomic DNA region near the HR sites, on both WT and mutant samples) and primer
SP35 (specific for the hygromycin B resistance gene, only present if the HR was successful). Reactions
D should result in the amplification of a DNMT3b gene fragment with 883 nt, in the lines where the gene
wasn’t replaced (WT genotype, Figure 26) and a fragment with 1289 nt, resulting from the annealing of
both SP33 and SP35 primers, on the Δdnmt3b lines. In the case of WT genotype lines, another band
could also be amplified (3942 nt) due to the annealing of SP30 and SP33 primers, although this band is
too big for the extension time used for the amplification reactions (Figure 26).
As observable from Figure 27, from both C and D reactions, the only fragments obtained had
similar sizes to the ones obtained from the WT tissue sample (positive control, Figure 27 reactions C
WT and D WT samples). The band detected in samples from reactions C has around 2100 nt (Figure
27, reactions C #1-10), while from reactions D the bands detected have between 800-900 nt (Figure 27,
reactions D #1-10).
In Figure S12, the individual in-tissue PCR reactions designed to amplify only one fragment per
reaction and to confirm the multiplex results are shown. Mix A contained primers SP31 and SP32 in
order to evaluate the presence of the 2137 nt DNMT3b gene-specific amplified fragment (Figure 26).
Reactions B (primers SP32 and SP34) and reactions C (SP33 and SP35) were designed to confirm the
correct replacement of the DNMT3b gene by the mCherry and hygromycin B resistance coding
sequences as well as the presence of the 5’ and 3’ flanking regions, respectively (Figure 26). From
reactions B, the expected band had 1292 nt while, in reactions C the size of the expected ampliied
53
fragment was 1289 nt. Finally, reactions D intended to access the presense of the hygromycin resistance
gene in the tissue DNA using primers CR2 and CR3, by amplifing a DNA fragment of 858 nt (Figure 26).
As seen in Figure S12, no amplification was detected in any of the reactions B samples. In
reactions C, the only observable band is from the reaction in which plasmid DNA was used (Figure S12
C, P well), but this band has over 1400 nt and not the 1289 nt expected. In reactions A, a band around
2100 nt was amplified in all the selection-surviving colonies as well as in the WT tissue, used as positive
control (Figure S12 A#1-10 and WT wells). The hygromycin resistance gene sequence appears to be,
at least in part, present in the tissue from colonies number 1, 3, 4, 5 and 10 based on the amplification
of a DNA fragment with sizes around 800-900 nt in these samples, as well as in the plasmid DNA (used
as positive control) (Figure S12 D#1, D#3-5, D#10 and D P wells). Amplification was never observed for
the negative control reactions, where reaction buffer was used as a replacement for template DNA –
containing tissue sample (Figure S12 B wells).
Figure 27: 1% agarose gel loaded with the in-tissue multiplex PCR reactions C and D, used for
genotyping the selection-surviving colonies from the transformation of Physcomitrella patens protoplasts
with the pSP3b plasmid. 1kb ladder (NEB) was used to estimate the size of amplified fragments. The size of the ladder’s bands are presented on the left part of the image. Wells numbered #1 to #10 represent the selection-surviving colonies number and the WT samples represent the reactions using wild-type tissue as template DNA - containing samples used as positive control for the reactions.
Antherozoids lack autofluorescence and can be labelled with FDA.
As described in the introduction the technique used for the collection of the antherozoid samples
used for the transcriptomic atlas involved dissecting the antheridia on the day of the antherozoid release
(15 days after sexual phase induction), placing them on water and collecting the released clusters of
antherozoids under a microscope using a micromanipulator, making it a very time-consuming process.
The first step to develop a more time-efficient way of collecting the antherozoids involved
observing antherozoid clusters (Figure 28 B and C) and isolated sperm cells (Figure 28 A), released
from manually dissected mature antheridia into a sperm-nutritive solution (Table 4) and assess their
54
approximate size and life-time in this solution. The approximate diameter of an antherozoid was of 10
µm (total antherozoid) and that of the antherozoid body (excluding the two flagella) of 5 µm (Figure 28
A). The antherozoids were found to be alive for a maximum of 1 h while in the sperm-nutritive solution
(Carlos Ramirez, oral communication) and in average they survived for around 45 min. These were also
observed in different channels (e.g. RFP and GFP channels), in order to confirm the previous
observations by Marcela Coronado (personal communication) that the antherozoids did not show
autofluorescence.
Figure 28: Bright field pictures of a single antherozoid (A) and clusters of antherozoids (B and C).
Antherozoids are released from P. patens mature antheridia, manually dissected into sperm nutritive solution. Mature antheridia are indicated by white arrows and the cluster of antherozoids shown on panel C is limited by a green dashed line. Scale bars = 10 μm.
With the goal of developing a more time-efficient method to collect P. patens antherozoids,
namely by the use of fluorescent-assisted cell sorting (FACS), we first had to label these cells. The
fluorescent labelling of the antherozoids was achieved by adding fluoresceín diacetate (FDA) to the
sperm-nutritive solution in which the mature antheridia were dissected and the antherozoids released.
Afterwards, the samples were observed under oil immersion at 100x magnification in order to detect if
releasing of the antherozoids clusters had occurred and if they were labelled. In Figure 29 A a labelled
isolated antherozoid in bright field is shown, its green fluorescence (Figure 29 B), present in both the
antherozoid body and its flagella, as well as the absence of autofluorescence, analysed by the red
fluorescence using the red channel (Figure 29 C). The merged picture of all three channels can be seen
in Figure 29 D.
Using the same method, labelled clusters were also observed and four examples with the merged
picture of the three channels used (bright field, green and red channels) can be seen on Figure 30. With
these observations, it was possible to perceive that the antherozoids in these clusters were labelled in
the green channel and showed no significant autofluorescent signal, while antheridia cells and tissue
55
debris showed a more bright red signal (chlorophyll autofluorescence) compared to the green (FDA)
signal (Figures 30 B and D).
Figure 29: FDA labelled antherozoid. Mature antheridia were manually dissected into sperm nutritive solution
supplemented with fluoresceín diacetate (FDA) and the antherozoids were released. Microscopic observation was achieved at 100x magnification under oil immersion. A: Widefield image (15 ms of exposure time); B: Green fluorescent signal detected (100 ms exposure time); C: Red fluorescent signal – autofluorescence, detected in the red mCherry fluorescent protein channel (250 ms exposure time); D: Merged image of the observed antherozoid in widefield, green and red channels. Scale bars = 10 μm.
Figure 30: Labelled antherozoid clusters from Physcomitrella patens. Mature antheridia were manually
dissected into sperm nutritive solution supplemented with fluoresceín diacetate and samples were observed at 100x amplification under oil immersion. Images from bright field (15 ms exposure), green fluorescence (100 ms exposure on GFP channel) and red fluorescence (200 ms exposure on RFP channel) were merged. Scale bars = 10 μm.
56
FACS sorting of antherozoids.
After successfully labelling of intact antherozoids with FDA, the next step in the development of a
time-efficient protocol to collect the antherozoids was to test, if sorting of these cells was possible by an
automated process, such as FACS.
In order to allow the samples analysis by flow cytometry, the particles in the sample could not be
bigger than 50 µm in diameter. Therefore, mature antheridia and antherozoids clusters had to be filtered
out from the sample before FACS. 10 µm mesh and 28 µm mesh filters were tested, and isolated
antherozoids were observed in the flow-through solutions obtained after the samples’ filtering with either
type of mesh (Figure 31 A, B – 10 µm mesh and C, D - 28 µm mesh). When the sperm-nutritive solution
was supplemented with FDA, the isolated antherozoids showed a bright green and no significant red
fluorescent signals, as can be seen on Figure 31 B (10 µm mesh) and D (28 µm mesh).
Figure 31: Isolated antherozoids obtained after antheridia sample filtering. Mature antheridia were manually dissected into sperm-nutritive solution in order to be filtered and obtain isolated antherozoids. All pictures were obtained by imaging the samples under oil immersion at 100x amplification. A: Bright field image (20 ms exposure) of an antherozoid observed from the solution filtered with a 10 µm mesh. B: Merged image of the bright field, green signal channel (GFP channel, 100 ms exposure) and red signal channel (RFP, 250 ms exposure) of the antherozoid observed on image A. C: Bright field image (20 ms exposure) of an antherozoid detected in the solution filtered with the 28 µm mesh. D: Merged image of the bright field, green signal (GFP channel, 100 ms exposure) and red signal (RFP, 250 ms exposure) of the antherozoid observed on image C. Scale bars = 10 µm.
Next, new samples were prepared into sperm-nutritive solution and analyzed in the MoFlo cell
sorter. After filtration, a portion of flow through solution was examined - unstained sample. Then, FDA
was added to the remaining sample - FDA stained sample. The same analysis performed on part of the
unstained sample was repeated for the stained sample. The results of the flow cytometric analyses are
shown in Figures 32 to 34, where panels A1-3 display the data from the unstained samples and panels
B1-3 show the results from the stained samples.
FDA+ polygon present on the panels numbered 1 identify the area considered to have high green
signals and a relative low red signal (FDA+ events). Polygons named Red and R2 (panels 2) represent
the area of the plot where the cells are considered to have a positive red signal (autofluorescent cells)
57
and the area where the cells are considered to be negative for both red and green signal, respectively.
Finally the panels numbered 3 show the count table obtained by counting all the events in the sample
(Total) and the FDA+ events.
Figure 32 presents the results obtained from a sample filtered with the 10 µm mesh. The
unstained sample (Figure 32 A1-3) shows the distribution of the red and green signals of the events in
the sample, the total amount of events (cells) analysed and the total FDA positive (FDA+) cells. Most of
the cells were considered to be negative for both signals (low RFP, FL4 and FL1 signals), some were
considered to have a high red signal (high RFP and FL4 signal) and with different degrees of green
signal intensity (FL1 log) (Figure 32, A1 and A2). From the 4648 total cells analysed, only 55 (~ 1 %)
were considered FDA+ cells due to their significant signal in the green channel (Figure 32, A3).
After the addition of FDA to the remaining part of the sample and its analysis (labelled sample -
Figure 32, B panels), most of the analysed cells were still considered negative for both red and green
signals (Figure 32, B1 and B2, R2 polygon) or red positive cells (Figure 32, B2 Red polygon). From the
total amount of events in the sample (33991), 3374 were considered to have a low red signal and a
positive green signal (FDA+ cells - Figure 32, B1 polygon FDA+), representing almost 10 % of the
sample. A significant value when compared to the 1 % from the unstained sample results. The FDA
positive cell population was sorted into a microscope slide and observed, but no antherozoids or any
other cell types were detected (data not shown).
Similar results are shown on Figure 33, but this time the 28 µm mesh was used to filter the
dissected mature antheridia sample and obtain the final cell suspension to be analysed. In the unstained
part of the sample (Figure 33, panels A), most of the cells were considered negative for both red and
green signals (Figure 33, A2 R2 polygon), some were considered to be only red labelled (Figure 33, A2
Red polygon) and almost none event was observed in the FDA+ polygon shown in Figure 33, A1. From
the 6480 events analysed in this sample, only 2 were considered as FDA+ events (0.03 %) (Figure 33,
A3).
Analysing the remaining sample after FDA addition (Figure 33, panels B) an increase in the FDA+
events (FDA+ polygon in Figure 33, B1 panel) was observed, although most remained negative for both
signals (Figure 33, B2 R2 polygon) or only red labelled (Figure 33, B2 Red polygon). From a total of
70421 events analysed 90 were considered as FDA+, corresponding to 0.13 % of the sample (Figure
33, B3). Panel C of Figure 33 shows the same plot as B1 panel but only showing the FDA+ events, and
thus the population of interest.
58
Figure 32: Flow cytometric analysis of the flow through obtained using a 10 µm mesh. Mature antheridia were dissected into sperm-nutritive solution and then filtered using a 10 µm mesh. Each red dot represents a single event detected. A: Unstained sample results. A1 – Plot considering the green (x-axis, FL1 log signal) and red signals detected (y-axis, RFP log signal). The population of events considered as FDA stained are plotted inside the FDA+ polygon, characterized by an elevated green signal and a low red signal. A2 – Plot of the green (x-axis, FL1 log) and Texas Red (TxRed) signals (y-axis, red signal, autofluorescence). The R2 polygon delimits the population of events considered negative for both green and red signal and the Red polygon marks the area where the autofluorescent events are expected to show up (high red and low green signals). A3 – Table summarizing the results obtained from the A1 plot, in which the region named FDA+ is related to the events plotted inside that polygon and the region named total is related with the total amount of events analysed; count column show the number of events detected in each region, %Hist is related to the percentage that each of the events of the regions represent from the plot and % All is related to the percentage that the events in each region represent from the total amount of events. B: Analysis of the FDA-stained portion of the sample. B1 – Plot obtained considering the green (x-axis) and red signals detected (y-axis) for each event. B2 – Green (x-axis) and TxRed signals (y-axis) of each event. B3 – Table summarizing the results obtained from the B1 plot.
59
Figure 33: Flow cytometric analysis of the 28 µm mesh filtered flow-through sample. Mature antheridia were dissected into sperm-nutritive solution and then filtered using a 28 µm mesh. Each red dot represents a single event detected. A: Unstained sample. A1 – Plot of the green (FL1 log signal) and red signals detected (RFP log signal) from each event. A2 – Green (FL1 log) and Texas Red (TxRed) signals of the events. A3 – Summary of the results from the A1 plot. B: FDA-stained portion of the sample. B1 – Green and red signals from each event. B2 –
Green signal intensity vs signal emitted on the TxRed channel of each event. B3 – Table with the results obtained from the B1 plot. C: B2 plot showing only the events considered as part of the population of interest, with an elevated green signal (FDA labelled) and low red signal (no autofluorescence), corresponding to possible isolated antherozoids.
60
Figure 34: Flow cytometric analysis of the 28 µm mesh filtered samples' flow-through. Mature antheridia
were dissected into sperm-nutritive solution and then filtered using a 28 µm mesh. Each red dot represents a single event detected. A: Unstained sample. A1 – Events’ green (FL1 log signal) and red signals detected (RFP log signal). A2 – Plot of the green and Texas Red (TxRed) signals (red signal, autofluorescence) of the events analysed. A3 – Table with the distribution of the events analysed in the A1 plot, in which the region named FDA+ is related to the events plotted inside that polygon and the region named “total” is related with the total amount of events analysed. B: FDA-labelled sample portion. B1 – Results from the green vs the red signal from each event analysed. B2 – Detection of the green and the TxRed signals from each event. B3 – Table with the results obtained from the B1 plot. C1: Distribution of the events detected, based on their green and orange (FL2 log signal) signals. The R8 polygon demarks the region where non-red and green events can be detected, and therefore representing the population of interest area. C2: Table with the results obtained from the C1 plot, wherein the number of events from each area (Total or R8 polygon) are displayed, as well as the percentage that they represent in the total amount of events detected.
61
Due to the higher number of cells in the 28 µm sample, this filter type was the one used in further
assays. Figure 34 shows the results of the last flow cytometry analysis conducted in this study. In the
unstained sample (Figure 34, panels A) most of the cells are negative for red and green signals (Figure
34, A2 R2 polygon), some are considered to be only red labelled (Figure 34, A2 Red polygon) and the
green positive and red negative events (FDA+ polygon, Figure 34, A1) - the events of interest - represent
1 out of 1519 total events counted (~ 0.07 %) (Figure 34, A3).
The results from the stained sample (Figure 34, panels B) show a significant intensification of the
events present in the FDA+ polygon in Figure 34, B1 panel, representing 0.42 % of the total sample
(307 / 72722 events). As in the previously described results, most of the events in this sample were still
considered negative for red and green signals (Figure 34, B2 R2 polygon) or only red positive and green
negative (Figure 35, B2 Red polygon). From panel C1 of Figure 35, a plot of the green signal (FL1 log,
x-axis) by the FL2 log signal (orange, y-axis), one can observe a second diagonal being formed by the
signals of the population of interest.
From this experiment, 289 events belonging to the population of interested (FDA green labelled
and no red, autofluorescent events) were sorted and observed at 100x amplification under the
microscope. We were able to observe green labelled, isolated antherozoids, examples of which are
shown in Figure 35, although some debris was also observed (Figure 35, right image).
Figure 35: Antherozoids sorted by FACS. Images obtained by microscopic observation of the sorted part of the population of interest, isolated by flow cytometric analysis of the sample examined in Figure 34. A total of 289 events were sorted. In each image isolated antherozoids are shown, and on the right image some debris can also be observed (red labelled). Scale bars = 5 µm.
62
Discussion
Cytosine methylation is a conserved feature in most eukaryotic genomes being catalysed by DNA
methyltransferases (DMTases). In the early land plant Physcomitrella patens, 7 genes potentially
encoding for DMTases were identified. MET1, CMT, DNMT2, DRM1, DRM2, DNMT3a and DNMT3b
have detectable homologs in P. patens genome (Table 1, Malik et al. 2012), but only the DRM2 and
DNMT3b genes appear to be significantly expressed in the antherozoids of this species (Table 2,
Hernández-Coronado, 2015; Ortiz-Ramírez et al.). DRM2 and DNMT3b DMTases are known de novo
methyltransferases from plants and animals, respectively (Cao et al., 2000; Hsieh, 1999) and previous
comparisons of 5-mC levels among 17 eukaryotic genomes revealed that P. patens had the highest
level of asymmetric CHH methylation (23.2 %) (Zemach et al., 2010).
Phylogenetic analysis of P. patens de novo methyltransferases, considering both the full protein
sequences and only the 5-mC methyltransferase domains, was used to evaluate the phylogenetic
position of these DMTases (Figures 10, S1 and S2). From these analyses it is possible to note that the
DRM1 and 2 sequences from P. patens always cluster together, with high bootstrap values ( > 80) and
closer to the remaining DRM sequences used (Figures 10 and S1), being consistent with their
identification as members of this protein family (Malik et al., 2012).
P. patens DNMT3a and DNMT3b cluster together in all phylogenetic trees obtained in this work,
with bootstrap values over 93 and closer to the other DNMT3 sequences (Figure 10 and S2 B), except
in the maximum likelihood tree obtained considering the full protein sequences of all DNMT3 and P.
patens DRM1 and DRM2 sequences as outgroup (Figure S2 A). In fact, on the tree shown on Figure
S2A, DRM1 and DRM2 sequences from P. patens cluster with the animal DNMT3 sequences and P.
patens DNMT3 sequences form the outgroup. This was not expected, however it can be explain due to
the observation that these two sequences (DNMT3a and DNMT3b) from P. patens show a particular
domain in their sequence - DUF3444, that is not found in any other known DMTase (Malik et al., 2012).
Another unexpected result from the phylogenetic analysis performed was the cluster formed by
DNMT1/MET1 sequences not representing an outgroup in the tree obtained from the alignment of the
full sequences (Figure 10 A). One possible explanation for this can be the fact that the algorithm will join
similar sequences by pairs and then compute the distance between that pair (now considered as only
one element) and all the other elements/groups. Consequently, since the DNMT1/MET1 cluster is
represented by 2 sequences from plants (A. thaliana and P. patens) and only one animal (human), when
the group is considered only as one element it may become more similar to the DRM cluster, since only
plant sequences are present there. Therefore, this may represent a consequence from the selection of
the DNMT1/MET1 sequences and the evolutionary position of the organisms to where they belong.
Overall, the bootstrap values obtained in all phylogenetic trees are not always high and do not
allow for a detailed analysis of all the clusters due to low confidence in the groups formed. Although in
all the trees, DNMT3a sequences group together between different animal species as the DNMT3b
sequences. Moreover, the clustering among the sequences of the same protein have a direct correlation
with evolution of the represented species: human (Homo sapiens) group with taurus (Bos taurus), then
mice (Mus musculus), chicken (Gallus gallus) and finally zebra fish (Danio rerio). The same is not
observed for the DRM sequences since DRM1 and DRM2 sequences do not form monophyletic groups
63
(Figures 10, S1 and S2). This can have several reasons, including the nomenclature of the proteins
themselves, different ploidy levels among species and the lower resolution and annotation of some plant
genomes. Despite some small differences, the trees considering the same set of sequences, either full
or the 5-mC methyltransferase domain sequence are very similar with the more striking difference being
the already described clustering of the DRM sequences of P. patens with the animal DNMT3 sequences
in the tree shown on Figure S2.
Bearing in mind the high levels of CHH methylation detected in P. patens, the presence of 4 genes
coding for possible de novo methyltransferases and the expression of only DRM2 and DNMT3b in the
antherozoids, we decided to study the function of the DRM2 gene in Physcomitrella patens. To that end,
two knockout lines (Δdrm2#1 and Δdrm2#2) were previously generated (by Anna Thamm and Marcela
Coronado) through the transformation of protoplasts with the pAT05 plasmid. Sequencing of the pAT05
plasmid was performed by NGS in this work, allowing us to obtain the complete sequence of the pAT05
plasmid (Figure 11) in a faster and more reliable (due to the high coverage obtained) method, when
compared to standard Sanger sequencing. During this work, genotyping of both Δdrm2#1 and Δdrm2#2
confirmed the deletion of DRM2 gene in these lines (Figures 13 and S3). Fertilization rates of WT and
Δdrm2 were accessed with the only significant difference detected being in the F0 between WT#1 and
Δdrm2#1 fertilization rates (p-value = 0.0006, Figure 14, Table S4). This difference was not detected
between the WT#2 and Δdrm2#2 probably due to high standard destination among the samples of these
lines (Figure 14, Table S4). The standard deviation of the samples are taken into account during t-test
analysis with the aim of conclude if the samples are different. The differences detected between
Δdrm2#1 and Δdrm2#2 lines can also be due to different insertion’s copy and/or due to different ploidy
levels (Schween et al., 2005). No differences were detected in the F1 and F2 generations, meaning that,
if any difference in the fertilization between WT and DRM2 deletion rates exists, it’s only in the F0
generation, being restored after the first fertilization event (Figure 14). This can be explained by several
different hypotheses. It is possible that the SCs that achieve fertilization are the ones able to cope with
DRM2 absence namely by the lack of TEs’ insertions on detrimental regions, and that, due to selection,
the next generations were no longer affected by the lack of DRM2 in the antherozoids. Another possible
explanation is that just after fertilization, the expression of DRM1 and/or DNMT3a, detected on both
archegonia and sporophyte development, could compensate for the potential hypomethylation of DRM2-
regulated regions. Alternatively, the expression of DNMT3b in the antherozoids may be sufficient to
compensate for the deletion of DRM2.
The fact that the WT values are slightly higher in the Δdrm2 lines may be explained if the
transformation process influences the fitness of the plants (fact also observed by Stefan Rensing,
University of Marburg, Germany, personal communication), despite not being statistically significant.
To further explore the hypothesis of DNMT3b compensating the DRM2 deletion in the
antherozoids, the pSP3b plasmid was cloned (Figure 9) and PEG mediated protoplast transformation
of the WT line was performed, with the purpose of obtaining Δdnmt3b lines. The transformation was
followed by two rounds of selection and only 14 colonies were obtained after the first round of selection,
since about half of the plates got heavily contaminations after the transformation protocol. Afterwards,
in-tissue multiplex (Figure 27) as well as individual PCR reactions (Figure S12) were used to genotype
64
the 10 colonies that survived the second round of selection, using a WT tissue sample as a positive
control for the WT genotype.
As it can be seen from Figure 28, as well as from Figure S12, the in-tissue PCR reactions with
the addition of PVP-40 (to precipitate the phenolic compounds present on the tissue samples) worked,
but in all the 10 surviving colonies only bands corresponding to WT genotype were observed. From the
individual PCR reactions D (Figure S12, D), wherein primers specific for the hygromycin resistance gene
were used, amplification of a fragment with a similar size to the obtained from the plasmid sample was
detected in the samples from colonies numbers 1, 3, 4, 5 and 10. This can indicate that, either
homologous recombination occurred in a different position in the genome or that the resistance gene is
still present on the tissue but not integrated on the genome, since P. patens cells can keep
extrachromosomal DNA for some time (Ann-Cathrin Lindner, IGC, personal communication).In order to
obtain and analyse the F1 generation of the WT and Δdrm2 lines, the germination of F0 spores was
required. After the sterilized spores were stored at 4 ºC during 14 weeks (3.5 months) and allowed to
germinate, all 21 days-old WT colonies showed a more irregular shape with a few gametophores
developing, probably trying to expand and looking for better conditions to grow (Figure 17) while colonies
of Δdrm2#2 line showed a more regular round shape and were smaller than the WT ones (Figures 18).
As for Δdrm2#1 line, no colonies were obtained (data not showed). This effect suggested some sort of
problem in spore germination and/or colony growth after cold storage of spores, with Δdrm2#1 spores
being highly affected (maybe losing viability) and Δdrm2#2 spores more affected than the WT’s.
This observation led us to design a more exhaustive experiment where spores would be
germinated immediately after sterilization (freshly sterilized spores) and every 2 weeks after being stored
at 4 ºC, until 14 weeks of storage, in order to try to assess possible differences in the growth rate and
final dry weight of the colonies growth from spores of different lines and after different periods of cold
storage. Total colony area (determined after 3, 5, 7, 10, 15 and 21 days of growth) had already been
used by Saavedra et al., (2011) to access protonema growth defects in mutants for enzymes involved
in lipid messenger synthesis and the and at 21 days of growth and whenever possible, 25 colonies dry
weight would be determined. Dry weight of petri-dishes with P. patens colonies germinated from spores
were reported in 2005 by Schween et al., but in both cases no details about the performed methodology
or analysis were described.
As expected, the colony area increased during the growth of the colonies (Figures 21 and 22)
and, although some statistically differences were observed between WT and Δdrm2’s areas, they do
not seem to indicate a clear difference between the lines since, in some samples the average colony
area of WT samples was higher and in others it was lower when compared to the respective Δdrm2 line
(Tables S5 and S8). ANOVA analysis of samples of the same genotype revealed some differences, but
as for the t-test they do not indicate a clear tendency of the colonies to grow differently according to the
time of cold storage of spores (Tables S6, S7, S9 and S10).
From the analysis of the colonies dry weight data from WT#1 and Δdrm2#1 lines (Figure 23 and
Table S11), no statistically significant differences were detected either by t-test or ANOVA analyses
performed (Figure 23, Tables S1, S13 and S14). This indicated that the cold storage of spores for
different periods of time does not seem to affect the 21 days old colonies’ dry weight from WT#1 and
65
Δdrm2#1 lines. From both the colonies area and their dry weight analyses, no biologically and consistent
relevant differences were detected between WT and Δdrm2 lines. Nevertheless, these methods can
now be used to study the protonema development in other lines.
The only consistent parameter observed from the data distribution plot of the dry weight of
samples from the WT#1 and Δdrm2#1 lines (Figure 23) and the plot of the WT#2 and Δdrm2#2 lines
(Figure 24), is the apparent homogenization of the samples dry weight (observed by the reduction of the
standard deviation values) after a certain time of cold storage of the spores. This is observable after 6
weeks of cold storage of WT#1 and Δdrm2#1 lines (Figure 23, Table S11) and after 8 weeks for WT#2
and Δdrm2#2 lines (Figure 24, Table 12). Such observation may be due to some sort of synchronization
or selection between the spores during the period of time storage such that the colonies obtained will
behave more homogenously. Synchronization could occur during the time when spores are kept
sterilized on water and the swelling of the spore is the first step of the germination process (Glime,
1983). In addition, freezing was found to be favourable for the germination of the spores of some
bryophyte species (During, 1979).
Despite the non-conclusive results from the colony area and dry weight measurements, smaller
colonies with a round shape were again observed during the progression of the detailed assay to study
colony growth performed, in samples from WT#1 after 6 weeks of cold storage of spores (3 / 10 plates)
and in colonies of Δdrm2#1 line after 10 weeks of storage of spores at 4 ºC (2 / 4) plates (Figure 19).
The appearance of such colonies on both WT and Δdrm2 samples indicates that this is not due to any
particular effect of the drm2 deletion. Due to the fact that the smaller and round colonies are always
detected on three plates and that one water bottle (with 9 mL of water) is used to spread the spores in
3 plates, the idea that the altered shape of the colonies could be due to differences in the water bottles
arose. All the bottles are always cleaned together and the same treatment is applied to all. Therefore
we decided to analyze the pH of the water in each bottle after sterilization.
The pH of the water of 10 different bottles was analyzed and it was found to vary between 6.9
and 8.9 (data not shown). pH affects many cellular processes, such as ionic exchanges, enzymatic
activities and compound solubility (Apinis, 1939). Therefore it could also have an effect on the
germination of the spores and growth of the colonies. In order to test this hypothesis, germination of
spores with water with pH values of 6.52, 6.9, 7.51, 7.8 and pH 8.93 was performed. We obtained
colonies in all samples, meaning that spores were able to germinate in the different pH values tested.
This is in accordance with Apinis (1939), who had already reported that spores from several bryophytes
could germinate in a wide range of pH values. No significant differences in colonies’ shape and aspect
were detected and no smaller, round colonies without gametophores were obtained (Figure 20),
indicating that water’s pH may not be the only reason influencing the phenotype of the colonies.
It is known that in bryophytes phytohormones may intervene in germination of spores (Shukla and
Kaul, 1991) protonema development and differentiation. Addition of exogenous auxins to the media
accelerates the transition from chloronema to caulonema during protonema growth of P. patens (Prigge
et al., 2010). Prigge and co-workers suggested the conservation of the auxin perception pathways
among land plants due to the presence, on P. patens genome, of four genes encoding known homologs
of AFB and three encoding homologs of IAA proteins, known to be involved in auxin signalling in A.
66
thaliana. In fact, silencing of the 4 AFB homologous P. patens genes as well as mutants for IAA
homologs, resulted in a suppression of caulonema differentiation originating colonies with appearance
similar to the small and round ones described in this work (Figures 18 and 19) (Prigge et al., 2010).
Small and round colonies, with defects on protonema growth and regeneration were also reported
in P. patens mutants for Dof1 transcription factor – a plant specific transcription factor known to be
involved in the regulation of stress responses, seed maturation and seed germination (Sugiyama et al.,
2012) and for RSL1/RSL2 transcription factor double mutants that were shown to be essential for the
differentiation of caulonema cells and to be positively regulated by auxin (Jang and Dolan, 2011). The
absence of caulonema cells in the protonema of PIPK1 and PIPK1 PIPK2 (the two P. patens’ genes
coding PIPKs) K.O. lines was one of the defects reported by Saavedra et al., in 2011. PIPKs are the
enzymes responsible for the synthesis of PtdIns-4, 5-bisphosphate, a eukaryotic lipid messenger with
important roles in cytoskeleton organization and intracellular vesicular trafficking, among others. Colony
area was decreased for Δpipk1 and Δpipk1-2 lines when compared to the WT area and the strong
reduction on caulonema cell growth was found to be related to defects in actin localization in these
mutants (Saavedra et al., 2011). Aspect of 2 weeks old colonies, growth on minimal media (KNOPS)
with cellophane disks, of Δrsl1-2 lines and their respective WT reported by Jang and Dolan (2011), can
be seen on Figure 36 (A and B), as well as 3 weeks old colonies (also grown on minimal media with
cellophane) of Δpipk1, Δpipk2 and Δpipk1-2 and respective WT lines reported by Saavedra et al., in
2011 (Figure 36, C - F).
The only study connecting cytosine methylation in Physcomitrella patens and alterations in the
growth of the protonema was reported by Dangwal et al., in 2014. Here, Δcmt lines displayed genome-
wide hypomethylation, with particular depletion of CHG methylation context and in CHG-methylation
rich loci a decrease in CHH methylation was also observed. Protonema proliferation of Δcmt plants was
arrested due to hindered chloronema growth and differentiation. The expression of stress related genes
was also found to be affected by CMT deletion in P. patens (Dangwal et al., 2014). During our work,
differences in the aspect of some P. patens colonies, wherein most of the colonies had irregular shapes
and developing gametophores by day 21 of growth while others exhibited a more round shape, were
smaller and had none, or a few, gametophores developing were detected among both WT and DRM2
deletion colonies. This seems to refute any link between the lack of DRM2-regulated 5-mC and the
growth and appearance of the 21 days-old colonies. Possible explanations for such observations may
be random effects during colony growth, oscillations in phytohormones, differences in the media of the
plates used, or some problem during material or sample preparation.
67
Figure 36: Examples of smaller, regular and round colonies lacking gametophores as reported for some
P. patens mutant lines. A and B: WT colony and colony of RSL/RSL2 double mutants (with arrested chloronema differentiation), grown in minimal media with cellophane for 2 weeks, reported by Jang and Dolan (2011); scale bar = 100 µm. C to F: 3 weeks old colonies from WT (C), Δpipk1 (D), Δpipk2 (E) and Δpipk1-2 (F), respectively, grown on minimal media with cellophane disks described by Saavedra and co-workers (2011); scale bar = 0.5 cm. Δpipk1 and Δpipk1-2 lines (C and F, respectively) show reduced colony area and the absence of caulonema cells in their protonema, when compared to the respective WT (C).
It is known that epigenetic reprogramming, involving the removal and de novo establishment of
5-mC marks on the genome, takes place during plant sexual reproduction (Calarco and Martienssen,
2011; Jullien et al., 2012) although, as to our knowledge, there are no reports about epigenetic
reprogramming in Physcomitrella patens to date. In general, the idea that the germline passively carries
the genetic information for the next generation is fading. Instead, the germline is becoming to be
considered a particularly active cell type that needs both to ensure genome viability (avoiding detrimental
lesions to be transmitted to the next generation) as well as to generate variation (e.g. by genetic
recombination) (Jablonka, 2013).
The antherozoids of Physcomitrella patens are a good example of a very specific cell type. They
are motile, biflagellated cells that are released in clusters from mature antheridia. The antherozoid
samples used by Marcela Coronado to complete the P. patens transcriptome atlas (Hernández-
Coronado, 2015) were collected using a very time-consuming method that involved the manual
dissection of mature antheridia to a microscope lamina, followed by the micromanipulation and collection
of individual antherozoid clusters. For further studies of these particular cells, namely by RNAseq or
bisulfite sequencing experiments, a time-efficient method to obtain antherozoid (e.g. by FACS) is
required.
Our first step in the development of such a method was to label the antherozoids and to confirm
the absence of autofluorescence, previously reported by Marcela Coronado in personal
communications. The labelling of P. patens antherozoids was achieved by the addition of fluoresceín
diacetate (FDA) to a sperm-nutritive solution (previously optimized by Carlos Ramirez, Table 4) followed
by the manual dissection of mature antheridia samples to the FDA-containing sperm-nutritive solution.
Green-labelled antherozoids were observed and the lack of autofluorescence was confirmed by imaging
on the RFP channel (Figures 29 and 30). With the aim of isolating antherozoid containing samples by
68
flow cytometry, manually dissected antheridia samples were filtered with 10 and 28 µm mesh. On
samples filtered with either mesh, individual and viable antherozoids were detected (Figure 31).
Subsequently, flow cytometric analyses of antherozoid containing samples were performed for
both the unstained and FDA-stained portions of the samples. In our first experiment, the 10 µm mesh
was used to filter the sample. Analysis of this sample allowed the identification of a population with the
characteristics of interest, representing 9.93 % of the total sample (Figure 32). This population was
sorted but no microscopic confirmation of the presence of antherozoids was possible, possibly since it
was already over 1 h after the filtering of the sample and the antherozoids are not known to survive
longer than 1 h in the sperm-nutritive solution (Carlos Ramirez, personal communication).
The remaining samples to be analyzed in the cell sorter were filtered with the 28 µm mesh and
less permissive signal limits were allowed, aiming to obtain a more distinct population of interest. This
was achieved by the decrease of the red signal intensity of the events to be considered of interest
(Figures 33 and 34). On our second analysis (Figure 33) only 0.13 % of the total amount of events were
considered to belong to our population of interest and on the final flow cytometric analysis (Figure 34)
0.42 % of the total events were considered as possible antherozoids.
From our final flow cytometric examination and using also detection of events’ orange
fluorescence (FL2 log signal), a second diagonal (highlighted by the R8 polygon) could be seen on the
C1 plot (Figure 34). This diagonal was constituted mostly by events belonging to the FDA+ events and
can therefore represent another signal to help in the detection of such events. Finally, from the 307
FDA+ events, we were able to sort 289 and confirm (by microscopic observation) the successful sorting
by FACS of isolated and viable (green labelled) antherozoids of the moss Physcomitrella patens (Figure
35). To our knowledge, this is the first report of an automated method that was successful in sorting P.
patens’ antherozoids.
In conclusion, we were unable to detect any consistent defect in the Δdrm2 lines analysed,
possibly due to DNMT3b compensation, whose deletion lines could not be obtained in the course of this
work. A link between the different phenotypes of the colonies described in this work, and the similar
reported phenotypes remains to be established. However, methods to analyse colony growth by
colonies area and dry weight, as well as genotyping by in tissue PCR were successfully implemented
during the progression of this work. Moreover, this work describes a novel automated method to
efficiently sort antherozoids of P. patens by FACS.
69
Future Perspectives
The moss Physcomitrella patens is an extant early land plants. Its genome is known to be heavily
methylated in all three sequence contexts. Opposed to vascular plants, P. patens 5-mC is particularly
enriched in repeated regions and absent in gene bodies, with extensive CHH methylation levels being
detected. This de novo methylation may be used to silence the high and dispersed amount of TEs found
on P. patens genome, a conserved feature among plants and vertebrates that reproduce sexually
(Zemach et al., 2010).
Four genes which code for de novo DNA methyltransferases: DRM1, DRM2 (homologous to
plant’s DRM proteins), DNMT3a and DNMT3b (with homology to de novo methyltransferases of animals)
were identified in P. patens genome (Malik et al., 2012). To our knowledge, no information about the
possible conservation of their functions along evolution is available, however reports showing a
somewhat conserved function between P. patens and other land plants’ MET1 and CMT were already
published (Dangwal et al., 2014; Noy-Malka et al., 2014; Yaari et al., 2015). A possible conservation of
functions of P. patens’ de novo DMTases will be studied further in the future.
During the analysis of two Δdrm2 lines no specific defects were detected, although differences
between both lines were observed. These differences can be due to different insertion copy numbers,
that will be assessed by quantitative real-time PCR (as described in Noy-Malka et al, 2013) and/or due
to different ploidy levels, to be determined by flow cytometry (as described in Schween et al., 2005).
Due to the possible compensation of DRM2 deletion by DNMT3b and the fact that no Δdnmt3b
lines were obtained in this study, both WT and Δdrm2 protoplasts will be transformed with the pSP3b
plasmid to obtain Δdnmt3b and Δdrm2 Δdnmt3b lines, respectively. Also, more Δdrm2 lines will be
required for more statistically significance of results. In-tissue multiplex and conventional PCR reactions
were used to genotype selection surviving colonies from regenerated protoplasts in this work. This
method worked, but since it was never attempted in our laboratory, DNA will be extracted from the
selection-surviving colonies and all the reactions repeated to confirm the results obtained.
With the intention of studying in detail the antherozoids of P. patens, we set out to develop an
automated method that would allow us to collect these cells in a time-efficient way and with sufficient
purity to perform molecular biology analysis. The FACS method developed allowed us to sort isolated
and viable antherozoids. This method needs further optimization to obtain a high number of isolated
antherozoids in a short amount of time. This can be achieved by decreasing the time required for the
preparation of each sample, either via reducing the time used for filtering steps or by avoiding the manual
dissection of the antheridia. However, one main concern is the purity of the sample, since for some
downstream analyses, such as RNAseq or bisulphite sequencing, the sample needs to be highly pure.
Purity of samples can be improved by adjusting the sorting parameters, such as the intensity of the
signals of the events to be sorted or the speed of sorting to allow for a more efficient separation.
In order to investigate cytosine methylation and its variation during Physcomitrella patens
reproduction, a general characterization of its methylome during different stages of its life cycle,
particularly in the gamete producing organs and the gametes themselves is needed. This will be
attempted via bisulphite sequencing of DNA samples isolated from different vegetative and reproductive
tissues, such as protonema, antheridia, antherozoids, archegonia and egg cells.
70
References
Apinis, A. (1939). “Data on the ecology of bryophytes III,” in Horti Botanica Universitatis Latviensis
Acta XI/XII:, 1–14.
Arif, M. A., Frank, W., and Khraiwesh, B. (2013). Role of RNA Interference (RNAi) in the Moss
Physcomitrella patens. Int. J. Mol. Sci. 14, 1516–40.
Auclair, G., Guibert, S., Bender, A., and Weber, M. (2014). Ontogeny of CpG island methylation and
specificity of DNMT3 methyltransferases during embryonic development in the mouse. Genome Biol.
15, 545.
Aufsatz, W., Mette, M. F., Van der Winden, J., Matzke, A. J. M., and Matzke, M. (2002). RNA-directed
DNA methylation in Arabidopsis. Proc. Natl. Acad. Sci. U. S. A. 99, 16499–506.
Bankevich, A., Nurk, S., Antipov, D., Gurevich, A. A., Dvorkin, M., Kulikov, A. S., et al. (2012).
SPAdes: a new genome assembly algorithm and its applications to single-cell sequencing. J. Comput.
Biol. 19, 455–77.
Baubec, T., Pecinka, A., Rozhon, W., and Mittelsten Scheid, O. (2009). Effective, homogeneous and
transient interference with cytosine methylation in plant genomic DNA by zebularine. Plant J. 57, 542–
554.
Bird, A. (2002). DNA methylation patterns and epigenetic memory. Genes Dev. 16, 6–21.
Böhmdorfer, G., Rowley, M. J., Kuciński, J., Zhu, Y., Amies, I., and Wierzbicki, A. T. (2014). RNA-
directed DNA methylation requires stepwise binding of silencing factors to long non-coding RNA. Plant
J. 79, 181–91.
Bond, D. M., and Baulcombe, D. C. (2014). Small RNAs and heritable epigenetic variation in plants.
Trends Cell Biol. 24, 100–107.
Bourc’his, D., and Bestor, T. H. (2004). Meiotic catastrophe and retrotransposon reactivation in male
germ cells lacking DNMT3L. Nature 431, 96–99.
Calarco, J. P., and Martienssen, R. a (2011). Genome reprogramming and small interfering RNA in
the Arabidopsis germline. Curr. Opin. Genet. Dev. 21, 134–9.
Cao, X., Aufsatz, W., Zilberman, D., Mette, M. F., Huang, M. S., Matzke, M., et al. (2003). Role of
the DRM and CMT3 Methyltransferases in RNA-Directed DNA Methylation. Curr. Biol. 13, 2212–2217.
Cao, X., and Jacobsen, S. E. (2002a). Locus-specific control of asymmetric and CpNpG methylation
by the DRM and CMT3 methyltransferase genes. Proc. Natl. Acad. Sci. U. S. A. 99, 16491–16498.
Cao, X., and Jacobsen, S. E. (2002b). Role of the Arabidopsis DRM methyltransferases in de novo
DNA methylation and gene silencing. Curr. Biol. 12, 1138–44.
Cao, X., Springer, N. M., Muszynski, M. G., Phillips, R. L., Kaeppler, S., and Jacobsen, S. E. (2000).
Conserved plant genes with similarity to mammalian de novo DNA methyltransferases. Proc. Natl. Acad.
Sci. U. S. A. 97, 4979–84.
Chen, C. C., Wang, K. Y., and Shen, C. K. J. (2012). The mammalian de novo DNA
methyltransferases DNMT3A and DNMT3B are also DNA 5-hydroxymethylcytosine
dehydroxymethylases. J. Biol. Chem. 287, 33116–33121.
Cho, S. H., Addo-Quaye, C., Coruh, C., Arif, M. A., Ma, Z., Frank, W., et al. (2008). Physcomitrella
patens DCL3 is required for 22-24 nt siRNA accumulation, suppression of retrotransposon-derived
71
transcripts, and normal development. PLoS Genet. 4, 1–13.
Coppedè, F., Bosco, P., Tannorella, P., Romano, C., Antonucci, I., Stuppia, L., et al. (2013). DNMT3B
promoter polymorphisms and maternal risk of birth of a child with Down syndrome. Hum. Reprod. 28,
545–550.
Coruh, C., Cho, S. H., Shahid, S., Liu, Q., Wierzbicki, A., and Axtell, M. J. (2015). Comprehensive
Annotation of Physcomitrella patens Small RNA Loci Reveals That the Heterochromatic Short Interfering
RNA Pathway Is Largely Conserved in Land Plants. Plant Cell 27, 2148–2162.
Cove, D. (2005). The moss Physcomitrella patens. Annu. Rev. Genet. 39, 339–358.
Cove, D. J., and Ashton, N. W. (1984). The hormonal regulation of gametophytic development in
bryophytes. Exp. Biol. Bryophyt., 177–201.
Dangwal, M., Kapoor, S., and Kapoor, M. (2014). The PpCMT chromomethylase affects cell growth
and interacts with the homolog of LIKE HETEROCHROMATIN PROTEIN 1 in the moss Physcomitrella
patens. Plant J. 77, 589–603.
During, H. J. (1979). Life Strategies of Bryophytes: A Preliminary Review on JSTOR. Lindbergia 5,
2–18.
Eamens, A., Vaistij, F. E., and Jones, L. (2008). NRPD1a and NRPD1b are required to maintain post-
transcriptional RNA silencing and RNA-directed DNA methylation in Arabidopsis. Plant J. 55, 596–606.
Feng, S., Cokus, S. J., Zhang, X., Chen, P.-Y., Bostick, M., Goll, M. G., et al. (2010a). Conservation
and divergence of methylation patterning in plants and animals. Proc. Natl. Acad. Sci. U. S. A. 107,
8689–8694.
Feng, S., Jacobsen, S. E., and Reik, W. (2010b). Epigenetic reprogramming in plant and animal
development. Science (80). 330, 622–627.
Finnegan, E. J., and Kovac, K. a (2000). Plant DNA methyltransferases. Plant Mol. Biol. 43, 189–
201.
Garcia, D., Garcia, S., Pontier, D., Marchais, A., Renou, J. P. P., Lagrange, T., et al. (2012). Ago
Hook and RNA Helicase Motifs Underpin Dual Roles for SDE3 in Antiviral Defense and Silencing of
Nonconserved Intergenic Regions. Mol. Cell 48, 109–120.
Glime, J. M. (1983). “Chapter 5-2: Ecophysiology of development: spore germination,” in Bryophyte
Ecology, 2–25.
Goll, M. G., Kirpekar, F., Maggert, K. a, Yoder, J. a, Hsieh, C.-L., Zhang, X., et al. (2006). Methylation
of tRNAAsp by the DNA methyltransferase homolog DNMT2. Science (80). 311, 395–398.
Hansen, R. S., Wijmenga, C., Luo, P., Stanek, a M., Canfield, T. K., Weemaes, C. M., et al. (1999).
The DNMT3B DNA methyltransferase gene is mutated in the ICF immunodeficiency syndrome. Proc.
Natl. Acad. Sci. U. S. A. 96, 14412–14417.
Henderson, I. R., and Jacobsen, S. E. (2007). Epigenetic inheritance in plants. Nature 447, 418–24.
Henikoff, S., and Comai, L. (1998). A DNA methyltransferase homolog with a chromodomain exists
in multiple polymorphic forms in Arabidopsis. Genetics 149, 307–318.
Hernández-Coronado, M. (2015). Comparative transcriptome analysis in the moss Physcomitrella
patens and the genetic basis of key reproductive innovations. phD thesis, Universidade Nova de Lisboa.
Portugal
72
Hsieh, C. L. (1999). In vivo activity of murine de novo methyltransferases, DNMT3a and DNMT3b.
Mol Cell Biol 19, 8211–8218.
Huang, Y., Kendall, T., Forsythe, E. S., Dorantes-Acosta, A., Li, S., Caballero-Perez, J., et al. (2015).
Ancient origin and recent innovations of RNA Polymerase IV and V. Mol. Biol. Evol.
Ichiyanagi, T., Ichiyanagi, K., Miyake, M., and Sasaki, H. (2013). Accumulation and loss of
asymmetric non-CpG methylation during male germ-cell development. Nucleic Acids Res. 41, 738–745.
Jablonka, E. (2013). Epigenetic inheritance and plasticity: The responsive germline. Prog. Biophys.
Mol. Biol. 111, 99–107.
Jaiswal, S. K., Sukla, K. K., Kumari, N., Lakhotia, A. R., Kumar, A., and Rai, A. K. (2015). Maternal
risk for Down syndrome and polymorphisms in the promoter region of the DNMT3B gene: A case-control
study. Birth Defects Res. Part A Clin. Mol. Teratol.
Jang, G., and Dolan, L. (2011). Auxin promotes the transition from chloronema to caulonema in moss
protonema by positively regulating PpRSL1 and PpRSL2 in Physcomitrella patens. New Phytol. 192,
319–327.
Jia, D., Jurkowska, R. Z., Zhang, X., Jeltsch, A., and Cheng, X. (2007). Structure of DNMT3a bound
to DNMT3L suggests a model for de novo DNA methylation. Nature 449, 248–51.
Jullien, P. E., Susaki, D., Yelagandula, R., Higashiyama, T., and Berger, F. (2012). DNA methylation
dynamics during sexual reproduction in Arabidopsis thaliana. Curr. Biol. 22, 1825–1830.
Kato, Y., Kaneda, M., Hata, K., Kumaki, K., Hisano, M., Kohara, Y., et al. (2007). Role of the DNMT3
family in de novo methylation of imprinted and repetitive sequences during male germ cell development
in the mouse. Hum. Mol. Genet. 16, 2272–2280.
Kawashima, T., and Berger, F. (2014). Epigenetic reprogramming in plant sexual reproduction. Nat.
Rev. Genet. 15, 613–624.
Kuhlmann, M., Finke, A., Mascher, M., and Mette, M. F. (2014). DNA methylation maintenance
consolidates RNA-directed DNA methylation and transcriptional gene silencing over generations in
Arabidopsis thaliana. Plant J.
Laird, P. W., and Jaenisch, R. (1996). The role of DNA methylation in cancer genetic and epigenetics.
Annu. Rev. Genet. 30, 441–464.
Landberg, K., Pederson, E. R. a, Viaene, T., Bozorg, B., Friml, J., Jönsson, H., et al. (2013). The
moss Physcomitrella patens reproductive organ development is highly organized, affected by the two
SHI/STY genes and by the level of active auxin in the SHI/STY expression domain. Plant Physiol. 162,
1406–19.
Law, J. A., and Jacobsen, S. E. (2011). Establising, maintaining and modifying DNA methylation
patterns in plants and animals. Nat Rev Genet. 11, 204–220.
Lister, R., Pelizzola, M., Dowen, R. H., Hawkins, R. D., Hon, G., Nery, J. R., et al. (2009). Human
DNA methylomes at base resolution show widespread epigenomic differences. Nature 462, 315–322.
Mahfouz, M. M. (2010). RNA-directed DNA methylation: mechanisms and functions. Plant Signal.
Behav. 5, 806–16.
Malik, G., Dangwal, M., Kapoor, S., and Kapoor, M. (2012). Role of DNA methylation in growth and
differentiation in Physcomitrella patens and characterization of cytosine DNA methyltransferases. FEBS
73
J. 279, 4081–94.
Manoharan, A., Roure, C. Du, Rolink, A. G., and Matthias, P. (2015). De novo DNA
Methyltransferases DNMT3a and DNMT3b regulate the onset of Igκ light chain rearrangement during
early B-cell development. Eur. J. Immunol.
Martienssen, R. a, and Colot, V. (2001). DNA methylation and epigenetic inheritance in plants and
filamentous fungi. Science (80). 293, 1070–4.
Matzke, M. a, Kanno, T., and Matzke, A. J. M. (2014). RNA-Directed DNA Methylation: The Evolution
of a Complex Epigenetic Pathway in Flowering Plants. Annu. Rev. Plant Biol., 1–25.
Mccue, A. D., Panda, K., Nuthikattu, S., Choudury, S. G., Thomas, E. N., and Slotkin, R. K. (2015).
ARGONAUTE 6 bridges transposable element mRNA-derived siRNAs to the establishment of DNA
methylation. EMBO J. 34, 20–35.
Morgan, H. D., Santos, F., Green, K., Dean, W., and Reik, W. (2005). Epigenetic reprogramming in
mammals. Hum. Mol. Genet. 14 Spec No, R47–58.
Mosquna, A., Katz, A., Decker, E. L., Rensing, S. A., Reski, R., and Ohad, N. (2009). Regulation of
stem cell maintenance by the Polycomb protein FIE has been conserved during land plant evolution.
Development 136, 2433–44.
Movahedi, A., Sun, W., Zhang, J., Wu, X., Mousavi, M., Mohammadi, K., et al. (2015). RNA-directed
DNA methylation in plants. Plant Cell Rep., 1–6.
Nam, E. J., Kim, K. H., Han, S. W., Cho, C. M., Lee, J., Park, J. Y., et al. (2010). The -283C/T
polymorphism of the DNMT3B gene influences the progression of joint destruction in rheumatoid
arthritis. Rheumatol. Int. 30, 1299–303.
Naumann, U., Daxinger, L., Kanno, T., Eun, C., Long, Q., Lorkovic, Z. J., et al. (2011). Genetic
evidence that DNA methyltransferase DRM2 has a direct catalytic role in RNA-directed DNA methylation
in Arabidopsis thaliana. Genetics 187, 977–9.
Nishiyama, T., Fujita, T., Shin-I, T., Seki, M., Nishide, H., Uchiyama, I., et al. (2003). Comparative
genomics of Physcomitrella patens gametophytic transcriptome and Arabidopsis thaliana: implication
for land plant evolution. Proc. Natl. Acad. Sci. U. S. A. 100, 8007–12.
Noy-Malka, C., Yaari, R., Itzhaki, R., Mosquna, A., Auerbach Gershovitz, N., Katz, A., et al. (2014).
A single CMT methyltransferase homolog is involved in CHG DNA methylation and development of
Physcomitrella patens. Plant Mol. Biol. 84, 719–35.
Okae, H., Chiba, H., Hiura, H., Hamada, H., and Sato, A. (2014). Genome-Wide Analysis of DNA
Methylation Dynamics during Early Human Development. PLoS Genet. 10, 1–12.
Okano, M., Bell, D. W., Haber, D. a, and Li, E. (1999). DNA methyltransferases DNMT3a and
DNMT3b are essential for de novo methylation and mammalian development. Cell 99, 247–57.
Ooi, S. K. T., and Bestor, T. H. (2008). The Colorful History of Active DNA Demethylation. Cell 133,
1145–1148.
Ortiz-Ramírez, C., Hernández-Coronado, M., Thamm, A., Catarino, B., Wang, M., Dolan, L., et al. A
Physcomitrella patens transcriptome atlas provides insights into the evolution and development of land
plants. Mol. plant, Submitted.
Paolillo, D. J. (1981). Swimming of Land Plants Sperms. Am. Inst. Biol. Sci. 31, 367–373.
74
Papa, C. M., Springer, N. M., Muszynski, M. G., Meeley, R., and Kaeppler, S. M. (2001). Maize
chromomethylase Zea methyltransferase2 is required for CpNpG methylation. Plant Cell 13, 1919–28.
Pélissier, T., Thalmeir, S., Kempe, D., Sänger, H. L., and Wassenegger, M. (1999). Heavy de novo
methylation at symmetrical and non-symmetrical sites is a hallmark of RNA-directed DNA methylation.
Nucleic Acids Res. 27, 1625–34.
Pires, N. D., and Dolan, L. (2012). Morphological evolution in land plants: new designs with old
genes. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 367, 508–18.
Pontier, D., Picart, C., Roudier, F., Garcia, D., Lahmy, S., Azevedo, J., et al. (2012). NERD, a Plant-
Specific GW Protein, Defines an Additional RNAi-Dependent Chromatin-Based Pathway in Arabidopsis.
Mol. Cell 48, 121–132.
Prigge, M. J., and Bezanilla, M. (2010). Evolutionary crossroads in developmental biology:
Physcomitrella patens. Development 137, 3535–43.
Prigge, M. J., Lavy, M., Ashton, N. W., and Estelle, M. (2010). Physcomitrella patens auxin-resistant
mutants affect conserved elements of an auxin-signaling pathway. Curr. Biol. 20, 1907–1912.
Rensing, S. a, Lang, D., Zimmer, A. D., Terry, A., Salamov, A., Shapiro, H., et al. (2008). The
Physcomitrella genome reveals evolutionary insights into the conquest of land by plants. Science (80).
319, 64–9.
Reski, R. (1998). Development, genetics and molecular biology of mosses. Bot. Acta 111, 1–15.
Rhee, I., Bachman, K. E., Park, B. H., Jair, K.-W., Yen, R.-W. C., Schuebel, K. E., et al. (2002).
DNMT1 and DNMT3b cooperate to silence genes in human cancer cells. Nature 416, 552–556.
Saavedra, L., Balbi, V., Lerche, J., Mikami, K., Heilmann, I., and Sommarin, M. (2011). PIPKs are
essential for rhizoid elongation and caulonemal cell development in the moss Physcomitrella patens.
Plant J. 67, 635–647.
Sasaki, H., and Matsui, Y. (2008). Epigenetic events in mammalian germ-cell development:
reprogramming and beyond. Nat. Rev. Genet. 9, 129–40.
Schaefer, and Zrÿd, J. (2001). The Moss Physcomitrella patens, Now and Then. Plant Physiol. 127.
Schumaker, K., and Dietrich, M. (1997). Programmed Changes in Form during Moss Development.
Plant Cell 9, 1099–1107.
Schween, G., Schulte, J., Reski, R., and Hohe, A. (2005). Effect of Ploidy Level on Growth,
Differentiation, and Morphology in Physcomitrella patens. Bryologist 108, 27–35.
Shukla, R. M., and Kaul, A. (1991). Influence of growth substances on spore germination of
Plagiochasma appendiculatum L. and L. Yushania. J. Bryol., 33–40.
Slotkin, R. K., Vaughn, M., Borges, F., Tanurdzić, M., Becker, J. D., Feijó, J. A., et al. (2009).
Epigenetic reprogramming and small RNA silencing of transposable elements in pollen. Cell 136, 461–
72.
Strotbek, C., Krinninger, S., and Frank, W. (2013). The moss Physcomitrella patens: methods and
tools from cultivation to targeted analysis of gene function. Int. J. Dev. Biol. 57, 553–64.
Stroud, H., Greenberg, M. V. C., Feng, S., Bernatavichute, Y. V, and Jacobsen, S. E. (2013).
Comprehensive analysis of silencing mutants reveals complex regulation of the Arabidopsis methylome.
Cell 152, 352–364.
75
Suetake, I., Miyazaki, J., Murakami, C., Takeshima, H., and Tajima, S. (2003). Distinct enzymatic
properties of recombinant mouse DNA methyltransferases DNMT3a and DNMT3b. J. Biochem. 133,
737–744.
Sugiyama, T., Ishida, T., Tabei, N., Shigyo, M., Konishi, M., Yoneyama, T., et al. (2012). Involvement
of PpDof1 transcriptional repressor in the nutrient condition-dependent growth control of protonemal
filaments in Physcomitrella patens. J. Exp. Bot. 63, 3185–3197.
Tiedemann, R. L., Putiri, E. L., Lee, J.-H., Hlady, R. A., Kashiwagi, K., Ordog, T., et al. (2014). Acute
Depletion Redefines the Division of Labor among DNA Methyltransferases in Methylating the Human
Genome. Cell Rep. 9, 1554–1566.
To, T. K., Saze, H., and Kakutani, T. (2015). DNA Methylation within Transcribed Regions. Plant
Physiol. 168, 1219–1225.
Viswanathan, C., and Jian-Kang, Z. (2011). RNA-directed DNA methylation and demethylation in
plants. Sci. China Press 52, 331–343.
Wassenegger, M., Heimes, S., Riedel, L., and Sänger, H. L. (1994). RNA-directed de novo
methylation of genomic sequences in plants. Cell 76, 567–576.
Watanabe, D., Suetake, I., Tada, T., and Tajima, S. (2002). Stage- and cell-specific expression of
DNMT3a and DNMT3b during embryogenesis. Mech. Dev. 118, 187–190.
Wolniak, S. M., Klink, V. P., Hart, P. E., and Tsai, C. W. (2000). Control of development and motility
in the spermatozoids of lower plants. Gravitational Sp. Biol. Bull. 13, 85–93.
Xie, S., Wang, Z., Okano, M., Nogami, M., Li, Y., He, W. W., et al. (1999). Cloning, expression and
chromosome locations of the human DNMT3 gene family. Gene 236, 87–95.
Xu, G. L., Bestor, T. H., Bourc’his, D., Hsieh, C. L., Tommerup, N., Bugge, M., et al. (1999).
Chromosome instability and immunodeficiency syndrome caused by mutations in a DNA
methyltransferase gene. Nature 402, 187–191.
Yaari, R., Noy-Malka, C., Wiedemann, G., Auerbach Gershovitz, N., Reski, R., Katz, A., et al. (2015).
DNA METHYLTRANSFERASE 1 is involved in mCG and mCCG DNA methylation and is essential for
sporophyte development in Physcomitrella patens. Plant Mol. Biol. 88, 387–400.
Zemach, A., Kim, M. Y., Hsieh, P. H., Coleman-Derr, D., Eshed-Williams, L., Thao, K., et al. (2013).
The Arabidopsis nucleosome remodeler DDM1 allows DNA methyltransferases to access H1-containing
heterochromatin. Cell 153, 193–205.
Zemach, A., McDaniel, I. E., Silva, P., and Zilberman, D. (2010). Genome-Wide Evolutionary
Analysis of Eukaryotic DNA Methylation. Science (80). 328, 916–9.
Zemach, A., and Zilberman, D. (2010). Evolution of eukaryotic DNA methylation and the pursuit of
safer sex. Curr. Biol. 20, 80–85.
Zhang, H., Chaudhury, A., and Wu, X. (2013). Imprinting in plants and its underlying mechanisms.
J. Genet. genomics 40, 239–47.
Zheng, B., Wang, Z., Li, S., Yu, B., Liu, J.-Y., and Chen, X. (2009). Intergenic transcription by RNA
polymerase II coordinates Pol IV and Pol V in siRNA-directed transcriptional gene silencing in
Arabidopsis. Genes Dev. 23, 2850–60.
Zheng, X., Pontes, O., Zhu, J., Miki, D., Zhang, F., Li, W., et al. (2008). ROS3, an RNA-binding
76
protein required for DNA demethylation in Arabidopsis. Nature 455, 1259–1262.
Zhong, X., Du, J., Hale, C. J., Gallego-Bartolome, J., Feng, S., Vashisht, A. a, et al. (2014). Molecular
mechanism of action of plant DRM de novo DNA methyltransferases. Cell 157, 1050–60.
Zhou, L., Cheng, X., Connolly, B. A., Dickman, M. J., Hurd, P. J., and Hornby, D. P. (2002).
Zebularine: A Novel DNA Methylation Inhibitor that Forms a Covalent Complex with DNA
Methyltransferases. J. Mol. Biol. 321, 591–599.
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Supplemental Material
Tables
Table S1: List of primers used in this work, their respective sequences (in 5’ to 3’ orientation) and their
purpose.
Primer name Primer sequence (5’ 3’) Purpose
AT 18 CTTTACGGTATCGCCGCTC
Δdrm2 genotyping
AT 23 ATGGAGGAGGAAGACACTTTGC
AT 24 ACGAAGCGAGCGATTGAGATAC
AT 25 GAACTTGTGGCCGTTTACGTC
AT 26 GGTAGGTTTGGAAGGATCAGGTC
AT 27 ATGCCTTCTTGGCTACACTTGG
CR 7 CGAGCTCGAATTCCCATGGA
CR 8 GCAAGGTGAGATGACGAGAGAT
dnmt3b_3KpnI_F GGAAGCGGTACCGGAAGAGAGCCTGAAACGTG Amplification of dnmt3b 3’region dnmt3b_3XbaI_R TAGCATTCTAGACTCATCTACTTCGTCCTTGGCA
dnmt3b _5GA_F TCACTATAGGGAATTTAAATTTAATTAAGCTTTCTTCGCCATCAATTCAAAGC Amplification of dnmt3b 5’ and mCherry sequences
dnmt3b_5GA_R CCATGGGCCCACTAGTTTAACGTAGCGTCACCAGTATCCTCT
cherry+T_GA_F TTAAACTAGTGGGCCCATGGTGAGCAAGGGCGAGG
cherry+T_GA_R TCTTTGATATTCTTGGAGGCGGCCGCAACGACGGCCAGTGAATTCC
pAT05_verify1 CAGCTATGACCATGATTACGAATTT
Confirmation of insert’s sequence
pAT05_verify2 TTCTTGGAGTAGACGAGAGTG
pAT05_verify3 ACCAAAATCCAGTACTAAAATCC
pAT05_verify4 CAGTGCCAAGCTAATTACCCTGT
pAT05_verify5 CTCGGAGGAGGCCATTG
pAT05_verify6 GTGAGTGGAACGAGCTTCGA
pAT05_verify7 CCTCCTTATCGAGCTCAT
SP 30 GCGAAGATTGAGGAGTTGAAGAG
Δdnmt3b genotyping
SP 31 CATCTACGCAATTGGTGACCG
SP 32 CATCAGTTCCACGGTTCCAGTC
SP 33 GTGCATAAAAGGTACTTGGCATC
SP 34 CCTTGATGATGGCCATGTTATCC
SP 35 GTACTCGCCGATAGTGGAAAC
CR 2 TGTAGGAGGGCGTGGATATG
CR 3 GCGAGTACTTCTACACAGCC
Table S2: List and respective composition of the solutions used in the protoplast transformation
protocol.
Solution name Solution composition
MMM 15 mM MgCl2; 1 % MES buffer (pH 5.6); 8.5 % D-Mannitol
35% PEG-4000 0.1 M Ca(NO3)2; 10 mM Tris buffer (pH 8.0); 35 % PEG-4000 (Merck); 7 % D-
Mannitol
Liquid proto KNOPS+GT media (Table 3); 0.36 M D-Mannitol
Top agar 8.5 % D-Mannitol; 1.4 % Agar (Sigma, A9799)
Proto plates KNOPS+GT media (Table 3); 0.36 M D-Mannitol; 10 mM CaCl2
- 2 -
Table S3: NCBI accession numbers for the non-Physcomitrella patens DNA methyltransferase protein
sequences used in our phylogenetic analysis.
Sequence identifier in the
phylogenetic trees
Sequence
accession number
Sequence identifier in the
phylogenetic trees
Sequence
accession number
DNMT1 (Homo sapiens) 12231019 ZMET3 (Zea mays) 212720705
MET1 (Arabidopsis thaliana) 332008394 DRM2 (Oryza sativa) 115450235
DRM1 (Arabidopsis thaliana) 257096638 DNMT3a (Homo sapiens) 166215081
DRM2 (Arabidopsis thaliana) 75184795 DNMT3b (Homo sapiens) 17375667
DRM3 (Arabidopsis thaliana) 18401465 DNMT3a (Bos taurus) 330417960
DRM1 (Glycine soja) 734312186 DNMT3b (Bos taurus) 164448558
DRM2 (Glycine soja) 734393438 DNMT3a (Mus musculus) 17374900
DRM1 (Aegilops tauschii) 475585873 DNMT3b (Mus musculus) 17374904
DRM2 (Aegilops tauschii) 475514122 DNMT3a (Gallus gallus) 82227308
DRM1 (Triticum urartu) 474156166 DNMT3b (Gallus gallus) 874507434
DRM2 (Triticum urartu) 474263464 DNMT3a (Danio rerio) 66392184
DRM2 (Arabis alpina) 674244855 DNMT3b (Danio rerio) 70887603
Table S4: Fertilization rates data. Obtained from F0, F1 and F2 generations of WT and Δdrm2 lines and the results (p-values) from the t-tests performed comparing WT and the respective Δdrm2 line. ns: non-significant; ***: significant difference detected.
Generation Line n Mean
fertilization rate Standard deviation
Standard error
t-test p-values
F0
WT#1 8 62.88 % 4.110 1.453 0.0006 (***)
Δdrm2#1 8 47.04 % 8.451 2.988 WT#2 8 62.78 % 9.035 3.194
0.1033 (ns) Δdrm2#2 8 51.46 % 14.98 5.296
F1
WT#1 5 49.20 % 8.260 3.690 1.0000 (ns)
Δdrm2#1 5 47.30 % 14.40 6.430 WT#2 5 53.00 % 9.110 4.070
0.7533 (ns) Δdrm2#2 5 50.20 % 18.70 8.350
F2
WT#1 5 64.00 % 13.30 5.960 0.8413 (ns)
Δdrm2#1 5 62.60 % 12.80 5.710 WT#2 5 61.20 % 14.70 6.570
0.8413 (ns) Δdrm2#2 5 56.90 % 12.10 5.420
- 3 -
Table S5: Colony area data of WT#1 and Δdrm2#1 lines as well was the t-test results of all the comparisons performed between lines. ND: not possible to determine (continues on next page).
Sample Days of growth
n Mean colony area (mm2)
Standard deviation
Standard error p-value of t-tests Sample Days of
growth n
Mean colony area (mm2)
Standard deviation
Standard error
WT#1 Fresh
3 ND ND
Δdrm2#1 Fresh
3 ND
5 13 0.02848 0.01821 0.00505 1.0000 (ns) 5 9 0.02792 0.01327 0.00442
7 42 0.10440 0.06819 0.01052 0.0121 (*) 7 9 0.17420 0.05193 0.01731
10 14 0.69800 0.32090 0.08576 0.0126 (*) 10 12 0.37570 0.26400 0.07620
15 17 5.54800 2.13600 0.51800 0.3973 (ns) 15 16 4.77800 3.14500 0.78620
21 18 20.870 15.420 3.63400 0.5616 (ns) 21 13 22.790 10.560 2.9300
WT#1 2w 4C
3 ND ND
Δdrm2#1 2w 4C
3 ND
5 ND ND 5 40 0.04504 0.01955 0.00309
7 48 0.16440 0.07907 0.01141 0.048 (*) 7 17 0.20100 0.05866 0.01423
10 50 0.87930 0.27920 0.03949 0.9215 (ns) 10 51 0.88560 0.23990 0.03359
15 58 6.37000 1.96000 0.25740 0.0353 (*) 15 45 7.29100 1.98700 0.29610
21 22 33.420 5.62300 1.19900 0.8148 (ns) 21 12 33.000 7.28900 2.10400
WT#1 4w 4C
3 ND ND
Δdrm2#1 4w 4C
3 11 0.02305 0.01445 0.00436
5 45 0.07852 0.03485 0.00520 0.7462 (ns) 5 33 0.07724 0.04111 0.00716
7 53 0.26930 0.07249 0.00996 0.1896 (ns) 7 45 0.24330 0.10240 0.01526
10 56 1.19100 0.26480 0.03539 0.0095 (**) 10 45 0.98200 0.43830 0.06534
15 44 9.17100 3.39200 0.51130 0.1481 (ns) 15 39 7.59300 3.35300 0.53680
21 22 35.070 7.95000 1.69500 0.0004 (***) 21 28 25.230 8.10800 1.53200
WT#1 6w 4C
3 ND ND
Δdrm2#1 6w 4C
3 3 0.02763 0.00500 0.00289
5 56 0.06818 0.02463 0.00329 0.8923 (ns) 5 60 0.07121 0.02812 0.00363
7 132 0.24790 0.09630 0.00838 0.0155 (*) 7 88 0.28160 0.07695 0.00820
10 98 1.24900 0.54230 0.05478 0.0052 (**) 10 66 1.51800 0.34990 0.04307
15 68 10.070 4.86300 0.58970 0.3375 (ns) 15 66 12.100 2.19200 0.26980
21 45 23.120 10.560 1.57500 0.2329 (ns) 21 40 27.160 5.26100 0.83190
- 4 -
Sample Days of growth
n Mean colony area (mm2)
Standard deviation
Standard error
p-value of t-tests Sample Days of growth
n Mean colony area (mm2)
Standard deviation
Standard error
WT#1 8w 4C
3 ND ND
Δdrm2#1 8w 4C
3 2 0.01293 0.00264 0.00187
5 31 0.04981 0.03322 0.00597 0.895 (ns) 5 36 0.04692 0.02743 0.00457
7 48 0.27650 0.62770 0.09060 0.8495 (ns) 7 47 0.19880 0.12080 0.01762
10 66 0.92290 0.49350 0.06075 0.635 (ns) 10 67 0.93780 0.65660 0.08022
15 64 4.54000 2.24800 0.28090 0.1047 (ns) 15 59 3.88800 2.50900 0.32670
21 31 28.370 8.65700 1.55500 0.4794 (ns) 21 16 25.500 8.35300 2.08800
WT#1 10w 4C
3 19 0.01869 0.00811 0.00186 0.0558 (ns)
Δdrm2#1 10w 4C
3 14 0.01372 0.01020 0.00273
5 100 0.09971 0.05307 0.00531 0.1992 (ns) 5 44 0.08622 0.04341 0.00655
7 103 0.48750 0.18920 0.01864 0.0001 (***) 7 54 0.34190 0.19540 0.02659
10 83 2.59500 0.76310 0.08376 0.0015 (**) 10 56 2.03900 1.12400 0.16570
15 21 14.630 2.78800 0.60840 0.0008 (***) 15 26 9.78600 5.18700 1.01700
21 13 35.870 5.39300 1.49600 0.0004 (***) 21 9 23.350 4.01000 1.33700
WT#1 12w 4C
3 6 0.02130 0.00345 0.00141 0.5686 (ns)
Δdrm2#1 12w 4C
3 13 0.02251 0.00986 0.00274
5 98 0.06465 0.02439 0.00246 < 0.0001 (****) 5 89 0.08745 0.03465 0.00367
7 59 0.24690 0.06730 0.00876 < 0.0001 (****) 7 65 0.30390 0.09015 0.01118
10 62 1.18000 0.23790 0.03021 0.0003 (***) 10 72 1.37100 0.35340 0.04165
15 35 7.68800 1.51900 0.25680 < 0.0001 (****) 15 40 10.100 1.92800 0.30480
21 23 19.680 4.50100 0.93850 < 0.0001 (****) 21 22 27.590 6.08200 1.29700
WT#1 14w 4C
3 ND ND
Δdrm2#1 14w 4C
3 ND
5 ND ND 5 11 0.03086 0.00825 0.00249
7 ND ND 7 14 0.18320 0.05453 0.01457
10 ND ND 10 ND
15 ND ND 15 8 4.47400 1.33700 0.47260
21 ND ND 21 10 17.790 5.58700 1.76700
- 5 -
Table S6: ANOVA statistical analysis of WT#1 colony area data. Colonies after 3 days of growth were analysed by t-test. ns: non-significant.
Day of growth Samples compared Statistical
result Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
3 10w vs 12w (t test) ns
7
Fresh vs 10w ***
10
Fresh vs 8w ns
15
Fresh vs 6w **
21
Fresh vs 4w **
5
Fresh vs 4w *** Fresh vs 12w *** Fresh vs 10w *** Fresh vs 8w ns Fresh vs 6w ns
Fresh vs 6w *** 2w vs 4w *** Fresh vs 12w * Fresh vs 10w *** Fresh vs 8w ns
Fresh vs 8w ns 2w vs 6w *** 2w vs 4w * Fresh vs 12w ns Fresh vs 10w **
Fresh vs 10w *** 2w vs 8w ns 2w vs 6w *** 2w vs 4w * Fresh vs 12w ns
Fresh vs 12w ** 2w vs 10w *** 2w vs 8w ns 2w vs 6w *** 2w vs 4w ns
4w vs 6w ns 2w vs 12w ** 2w vs 10w *** 2w vs 8w ns 2w vs 6w **
4w vs 8w ** 4w vs 6w ns 2w vs 12w * 2w vs 10w *** 2w vs 8w ns
4w vs 10w ns 4w vs 8w * 4w vs 6w ns 2w vs 12w ns 2w vs 10w ns
4w vs 12w ns 4w vs 10w *** 4w vs 8w * 4w vs 6w ns 2w vs 12w ***
6w vs 8w ns 4w vs 12w ns 4w vs 10w *** 4w vs 8w *** 4w vs 6w **
6w vs 10w ** 6w vs 8w ns 4w vs 12w ns 4w vs 10w ** 4w vs 8w ns
6w vs 12w ns 6w vs 10w *** 6w vs 8w *** 4w vs 12w ns 4w vs 10w ns
8w vs 10w *** 6w vs 12w ns 6w vs 10w *** 6w vs 8w *** 4w vs 12w ***
8w vs 12w ns 8w vs 10w *** 6w vs 12w ns 6w vs 10w ** 6w vs 8w ns
10w vs 12w *** 8w vs 12w ns 8w vs 10w *** 6w vs 12w ns 6w vs 10w **
7
Fresh vs 2w ns 10w vs 12w *** 8w vs 12w ns 8w vs 10w *** 6w vs 12w ns
Fresh vs 4w ***
10
Fresh vs 2w ns 10w vs 12w *** 8w vs 12w *** 8w vs 10w ns
Fresh vs 6w *** Fresh vs 4w * 15
Fresh vs 2w ns 10w vs 12w *** 8w vs 12w *
Fresh vs 8w * Fresh vs 6w ** Fresh vs 4w * 21 Fresh vs 2w ** 10w vs 12w ***
- 6 -
Table S7: ANOVA statistical analysis of Δdrm2#1 colony area data. ns: non-significant.
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
3
4w vs 6w ns
5
4w vs 6w ns
7
2w vs 6w ns
10
Fresh vs 8w ns
15
Fresh vs 14w ns
21
2w vs 4w ns
4w vs 8w ns 4w vs 8w * 2w vs 8w ns Fresh vs 10w *** 2w vs 6w *** 2w vs 6w ns
4w vs 10w ns 4w vs 10w ns 2w vs 10w ** Fresh vs 12w *** 2w vs 8w ** 2w vs 8w ns
4w vs 12w ns 4w vs 12w ns 2w vs 12w ** 2w vs 4w ns 2w vs 10w ns 2w vs 10w ns
6w vs 8w ns 4w vs 14w *** 2w vs 14w ns 2w vs 6w *** 2w vs 12w * 2w vs 12w ns
6w vs 10w ns 6w vs 8w ** 4w vs 6w ns 2w vs 8w ns 2w vs 14w ns 2w vs 14w ***
6w vs 12w ns 6w vs 10w ns 4w vs 8w ns 2w vs 10w *** 6w vs 8w *** 4w vs 6w ns
8w vs 10w ns 6w vs 12w ns 4w vs 10w ns 2w vs 12w *** 6w vs 10w ns 4w vs 8w ns
8w vs 12w ns 6w vs 14w *** 4w vs 12w ns 4w vs 6w *** 6w vs 12w ns 4w vs 10w ns
10w vs 12w ns 8w vs 10w *** 4w vs 14w ns 4w vs 8w ns 6w vs 14w *** 4w vs 12w ns
5
Fresh vs 2w ns 8w vs 12w *** 6w vs 8w *** 4w vs 10w *** 8w vs 10w *** 4w vs 14w ns
Fresh vs 4w ** 8w vs 14w ns 6w vs 10w ns 4w vs 12w ** 8w vs 12w *** 6w vs 8w ns
Fresh vs 6w ** 10w vs 12w ns 6w vs 12w ns 6w vs 8w *** 8w vs 14w ns 6w vs 10w ns
Fresh vs 8w ns 10w vs 14w *** 6w vs 14w * 6w vs 10w ns 10w vs 12w ns 6w vs 12w ns
Fresh vs 10w *** 12w vs 14w *** 8w vs 10w *** 6w vs 12w ns 10w vs 14w * 6w vs 14w *
Fresh vs 12w ***
7
Fresh vs 2w ns 8w vs 12w *** 8w vs 10w *** 12w vs 14w ** 8w vs 10w ns
Fresh vs 14w ns Fresh vs 4w ns 8w vs 14w ns 8w vs 12w ***
21
Fresh vs 2w ns 8w vs 12w ns
2w vs 4w ** Fresh vs 6w * 10w vs 12w ns 10w vs 12w ns Fresh vs 4w ns 8w vs 14w ns
2w vs 6w ** Fresh vs 8w ns 10w vs 14w ***
15
Fresh vs 2w ns Fresh vs 6w ns 10w vs 12w ns
2w vs 8w ns Fresh vs 10w ** 12w vs 14w ** Fresh vs 6w *** Fresh vs 8w ns 10w vs 14w ns
2w vs 10w *** Fresh vs 12w **
10
Fresh vs 2w ns Fresh vs 8w ns Fresh vs 10w ns 12w vs 14w *
2w vs 12w *** Fresh vs 14w ns Fresh vs 4w * Fresh vs 10w ** Fresh vs 12w ns
2w vs 14w ns 2w vs 4w ns Fresh vs 6w *** Fresh vs 12w *** Fresh vs 14w ns
- 7 -
Table S8: Colony area data of WT#2 and Δdrm2#2 lines as well was the t-test results of all the comparisons performed between lines. ND: not possible to determine; ns: non-significant (continues on next page).
Sample Days of growth
n Mean colony area (mm2)
Standard deviation
Standard error p-value of t-tests Sample Days of
growth n
Mean colony area (mm2)
Standard deviation
Standard error
WT#1 Fresh
3 2 0.00098 0.00032 0.00023 ND
Δdrm2#1 Fresh
3 ND
5 10 0.02443 0.02077 0.00657 0.5495 (ns) 5 11 0.02330 0.01223 0.00369
7 55 0.05724 0.02989 0.00403 < 0.0001 (****) 7 35 0.09810 0.03065 0.00518
10 75 0.28680 0.11330 0.01308 < 0.0001 (****) 10 60 0.52500 0.17450 0.02253
15 45 2.70600 0.95690 0.14260 < 0.0001 (****) 15 23 5.31900 1.78800 0.37270
21 40 13.9800 4.28700 0.67780 < 0.0001 (****) 21 20 21.8400 4.76800 1.06600
WT#1 2w 4C
3 ND ND
Δdrm2#1 2w 4C
3 ND
5 37 0.04173 0.01413 0.00232 < 0.0001 (****) 5 40 0.07563 0.03040 0.00481
7 88 0.10320 0.04480 0.00478 < 0.0001 (****) 7 84 0.25160 0.07103 0.00775
10 84 0.53760 0.20840 0.02274 < 0.0001 (****) 10 90 1.20500 0.26750 0.02820
15 58 2.49700 1.06300 0.13960 < 0.0001 (****) 15 28 6.42900 1.65800 0.31330
21 27 7.90500 2.03900 0.39240 < 0.0001 (****) 21 20 16.8800 5.07400 1.13500
WT#1 4w 4C
3 ND ND
Δdrm2#1 4w 4C
3 ND
5 10 0.03130 0.01190 0.00376 < 0.0001 (****) 5 27 0.06810 0.02096 0.00403
7 14 0.05184 0.02588 0.00692 < 0.0001 (****) 7 42 0.24120 0.09803 0.01513
10 19 0.17390 0.11450 0.02627 < 0.0001 (****) 10 39 1.12100 0.43690 0.06996
15 ND ND 15 33 5.55100 2.15100 0.37450
21 ND ND 21 29 14.3100 3.71600 0.69010
WT#1 6w 4C
3 ND ND
Δdrm2#1 6w 4C
3 ND
5 23 0.04426 0.01508 0.00314 0.1221 (ns) 5 20 0.03671 0.00920 0.00206
7 31 0.12540 0.03818 0.00686 0.2203 (ns) 7 26 0.13810 0.04085 0.00801
10 36 0.75850 0.28090 0.04681 0.0971 (ns) 10 41 0.66370 0.18050 0.02818
15 13 7.99200 2.97700 0.82560 < 0.0001 (****) 15 36 4.28700 0.75820 0.12640
21 13 16.8200 4.41300 1.22400 0.0105 (*) 21 29 13.2000 2.26100 0.41990
- 8 -
Sample Days of growth
n Mean colony area (mm2)
Standard deviation
Standard error
p-value of t-tests Sample Days of growth
n Mean colony area (mm2)
Standard deviation
Standard error
WT#1 8w 4C
3 2 0.04229 0.02108 0.01491 ND
Δdrm2#1 8w 4C
3 18 0.01749 0.01115 0.00263
5 34 0.07072 0.02493 0.00428 0.005 (**) 5 55 0.09521 0.03913 0.00528
7 52 0.25490 0.05998 0.00832 0.0002 (***) 7 63 0.31020 0.07506 0.00946
10 51 1.10500 0.31360 0.04391 0.3149 (ns) 10 61 1.05100 0.20650 0.02644
15 22 8.71500 4.10100 0.87430 0.026 (*) 15 32 6.39300 1.87500 0.33150
21 10 18.8900 8.84500 2.79700 0.5956 (ns) 21 29 19.2400 5.74100 1.06600
WT#1 10w 4C
3 ND ND
Δdrm2#1 10w 4C
3 3 0.01266 0.00311 0.00139
5 ND ND 5 5 0.08982 0.03214 0.00402
7 ND ND 7 7 0.33970 0.07555 0.00910
10 ND ND 10 10 1.49300 0.31450 0.04362
15 ND ND 15 15 9.21900 1.58600 0.23640
21 ND ND 21 21 22.4000 5.63500 1.12700
WT#1 12w 4C
3 ND ND
Δdrm2#1 12w 4C
3 3 0.00949 0.00175 0.00124
5 ND ND 5 5 0.07315 0.02219 0.00641
7 ND ND 7 7 0.27940 0.10190 0.02722
10 ND ND 10 10 1.09300 0.38440 0.10270
15 ND ND 15 15 2.64900 1.15300 0.36470
21 ND ND 21 21 14.1000 7.18000 1.53100
WT#1 14w 4C
3 25 0.01695 0.00677 0.00135 0.8535 (ns)
Δdrm2#1 14w 4C
3 13 0.01664 0.00427 0.00119
5 81 0.08773 0.03592 0.00399 < 0.0001 (****) 5 56 0.12180 0.04440 0.00593
7 88 0.40720 0.13190 0.01406 < 0.0001 (****) 7 57 0.50200 0.14680 0.01944
10 79 2.02400 0.55890 0.06288 < 0.0001 (****) 10 44 2.54500 0.76600 0.11550
15 30 12.2400 3.17200 0.57910 0.2039 (ns) 15 27 11.0500 3.88600 0.74780
21 11 31.9000 6.07900 1.83300 0.0006 (***) 21 16 23.6400 3.80400 0.95110
- 9 -
Table S9: ANOVA statistical analysis of WT#2 colony area data. ns: non-significant.
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
3
Fresh vs 8w ** 5 8w vs 14w ns
10
Fresh vs 4w ns
21
Fresh vs 14w ***
Fresh vs 14w ns
7
Fresh vs 2w ** Fresh vs 6w *** 2w vs 6w ***
8w vs 14w ns Fresh vs 4w ns Fresh vs 8w *** 2w vs 8w ***
5
Fresh vs 2w ns Fresh vs 6w ** Fresh vs 14w *** 2w vs 14w ***
Fresh vs 4w ns Fresh vs 8w *** 2w vs 4w ** 6w vs 8w ns
Fresh vs 6w ns Fresh vs 14w *** 2w vs 6w ns 6w vs 14w ns
Fresh vs 8w *** 2w vs 4w ns 2w vs 8w *** 8w vs 14w ns
Fresh vs 14w *** 2w vs 6w ns 2w vs 14w *** Fresh vs 6w ns
2w vs 4w ns 2w vs 8w *** 4w vs 6w *** Fresh vs 8w ns
2w vs 6w ns 2w vs 14w *** 4w vs 8w *** Fresh vs 14w ***
2w vs 8w *** 4w vs 6w ns 4w vs 14w *** 2w vs 6w ***
2w vs 14w *** 4w vs 8w *** 6w vs 8w ns 2w vs 8w ***
4w vs 6w ns 4w vs 14w *** 6w vs 14w *** 2w vs 14w ***
4w vs 8w *** 6w vs 8w *** 8w vs 14w ** 6w vs 8w ns
4w vs 14w *** 6w vs 14w ***
15
Fresh vs 2w ns 6w vs 14w ns
6w vs 8w ** 8w vs 14w * Fresh vs 6w *** 8w vs 14w ns
6w vs 14w *** 10 Fresh vs 2w *** Fresh vs 8w ***
- 10 -
Table S10: ANOVA statistical analysis of Δdrm2#2 colony area data. ns: non-significant.
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
Day of growth
Samples compared
Statistical result
3
8 vs 10w ns
5
6 vs 10w ***
7
4 vs 12w ns
10
4 vs 6w ***
15
2 vs 10w **
21
2 vs 4w ns
8 vs 12w ns 6 vs 12w ns 4 vs 14w *** 4 vs 8w ns 2 vs 12w ** 2 vs 6w ns
8 vs 14w ns 6 vs 14w *** 6 vs 8w *** 4 vs 10w * 2 vs 14w *** 2 vs 8w ns
10 vs 12w ns 8 vs 10w ns 6 vs 10w *** 4 vs 12w ns 4 vs 6w ns 2 vs 10w ns
10 vs 14w ns 8 vs 12w ns 6 vs 12w ** 4 vs 14w *** 4 vs 8w ns 2 vs 12w ns
12 vs 14w ns 8 vs 14w ns 6 vs 14w *** 6 vs 8w *** 4 vs 10w *** 2 vs 14w *
5
Fresh vs 2w ** 10 vs 12w ns 8 vs 10w ns 6 vs 10w *** 4 vs 12w ns 4 vs 6w ns
Fresh vs 4w * 10 vs 14w * 8 vs 12w ns 6 vs 12w * 4 vs 14w *** 4 vs 8w ns
Fresh vs 6w ns 12 vs 14w * 8 vs 14w *** 6 vs 14w *** 6 vs 8w ** 4 vs 10w ***
Fresh vs 8w ***
7
Fresh vs 2w *** 10 vs 12w ns 8 vs 10w *** 6 vs 10w *** 4 vs 12w ns
Fresh vs 10w *** Fresh vs 4w *** 10 vs 14w ** 8 vs 12w ns 6 vs 12w ns 4 vs 14w ***
Fresh vs 12w * Fresh vs 6w ns 12 vs 14w ** 8 vs 14w *** 6 vs 14w *** 6 vs 8w **
Fresh vs 14w *** Fresh vs 8w ***
10
Fresh vs 2w *** 10 vs 12w ns 8 vs 10w *** 6 vs 10w ***
2 vs 4w ns Fresh vs 10w *** Fresh vs 4w *** 10 vs 14w ns 8 vs 12w ** 6 vs 12w ns
2 vs 6w *** Fresh vs 12w *** Fresh vs 6w ns 12 vs 14w *** 8 vs 14w *** 6 vs 14w ***
2 vs 8w ns Fresh vs 14w *** Fresh vs 8w ***
15
Fresh vs 2w ns 10 vs 12w *** 8 vs 10w ns
2 vs 10w ns 2 vs 4w ns Fresh vs 10w *** Fresh vs 4w ns 10 vs 14w ns 8 vs 12w ns
2 vs 12w ns 2 vs 6w ** Fresh vs 12w *** Fresh vs 6w ns 12 vs 14w *** 8 vs 14w ns
2 vs 14w *** 2 vs 8w * Fresh vs 14w *** Fresh vs 8w ns
21
Fresh vs 2w ns 10 vs 12w ***
4 vs 6w * 2 vs 10w *** 2 vs 4w ns Fresh vs 10w *** Fresh vs 4w *** 10 vs 14w ns
4 vs 8w ns 2 vs 12w ns 2 vs 6w *** Fresh vs 12w ns Fresh vs 6w *** 12 vs 14w ***
4 vs 10w ns 2 vs 14w *** 2 vs 8w ns Fresh vs 14w *** Fresh vs 8w ns
4 vs 12w ns 4 vs 6w * 2 vs 10w * 2 vs 4w ns Fresh vs 10w ns
4 vs 14w *** 4 vs 8w ns 2 vs 12w ns 2 vs 6w ** Fresh vs 12w **
6 vs 8w *** 4 vs 10w *** 2 vs 14w *** 2 vs 8w ns Fresh vs 14w ns
- 11 -
Table S11: Dry weight data of WT#1 and Δdrm2#1, as well was the t-test results of the comparisons performed between these lines. ND: not possible to determine; ns: non-significant
Line Storage time (4 ºC)
n Mean of the colonies dry weight (mg)
Standard deviation
Standard error p-values of t-test Line Storage time
(4 ºC) n
Mean of the colonies dry weight (mg)
Standard deviation
Standard error
WT#1
0w 25 6.128 1.817 0.3634 0.9845 (ns)
Δdrm2#1
0w 25 5.968 3.562 0.7125
2w 25 5.44 2.797 0.5594 0.8690 (ns) 2w 25 5.296 3.256 0.6511
4w 25 6.344 2.783 0.5565 0.5604 (ns) 4w 25 6.496 3.018 0.6037
6w 25 6.996 1.102 0.2203 0.4783 (ns) 6w 25 7.068 0.7993 0.1599
8w 25 6.700 1.236 0.2472 0.2400 (ns) 8w 25 6.300 1.263 0.2527
10w 22 6.568 1.021 0.2177 0.9097 (ns) 10w 17 6.565 1.053 0.2554
12w 25 6.028 1.334 0.2669 0.7267 (ns) 12w 25 5.7400 1.590 0.3180
14w ND ND 14w 13 6.577 1.164 0.3229
Table S12: Dry weight data of WT#2 and Δdrm2#2, as well was the t-test results of the comparisons performed between these lines. ND: not possible to determine.
Line Storage time (4 ºC)
n Mean of the colonies dry weight (mg)
Standard deviation
Standard error p-values of t-test Line Storage time
(4 ºC) n
Mean of the colonies dry weight (mg)
Standard deviation
Standard error
WT#2
0w 25 7.788 3.086 0.6173 0.3984 (ns)
Δdrm2#2
0w 25 8.316 2.914 0.5828
2w 25 8.52 2.441 0.4881 0.3416 (ns) 2w 25 7.872 2.937 0.5875
4w 10 10.48 2.286 0.7229 0.0795 (ns) 4w 25 9.104 2.989 0.5978
6w 20 5.505 2.962 0.6624 0.9818 (ns) 6w 25 5.716 3.606 0.7213
8w 25 5.2 1.862 0.3724 0.0012 (**) 8w 25 6.772 1.177 0.2355
10w ND ND 10w 25 7.752 0.7287 0.1457
12w ND ND 12w 24 6.996 1.16 0.2368
14w 25 6.212 1.053 0.2106 < 0.0001 (****) 14w 25 7.856 1.432 0.2863
- 12 -
Table S13: ANOVA statistical analysis of WT#1 colony dry weight data. No colonies were obtained from spores stored at 4 ºC for 14 weeks. Therefore this sample was not used in statistical analysis. ns: non-significant.
Fresh 2w 4C 4w 4C 6w 4C 8w 4C 10w 4C
2w 4C ns . . . . .
4w 4C ns ns . . . .
6w 4C ns ns ns . . .
8w 4C ns ns ns ns . .
10w 4C ns ns ns ns ns .
12w 4C ns ns ns ns ns ns
Table S14: ANOVA statistical analysis of Δdrm2#1 colony dry weight data. ns: non-significant.
Fresh 2w 4C 4w 4C 6w 4C 8w 4C 10w 4C 12w 4C
2w 4C ns . . . . . .
4w 4C ns ns . . . . .
6w 4C ns ns ns . . . .
8w 4C ns ns ns ns . . .
10w 4C ns ns ns ns ns . .
12w 4C ns ns ns ns ns ns .
14w 14C ns ns ns ns ns ns ns
Table S15: ANOVA statistical analysis of WT#2 colony dry weight data. No colonies were obtained from spores stored at 4 ºC for 10 and 12 weeks therefore, this sample was not used in statistical analysis. ns: non-significant.
Fresh 2w 4C 4w 4C 6w 4C 8w 4C
2w 4C ns . . . .
4w 4C ns ns . . .
6w 4C ns * ** . .
8w 4C ns *** *** ns .
14w 14C ns ns ** ns ns
Table S16: ANOVA statistical analysis of Δdrm2#2 colony dry weight data. ns: non-significant.
Fresh 2w 4C 4w 4C 6w 4C 8w 4C 10w 4C 12w 4C
2w 4C ns . . . . . .
4w 4C ns ns . . . . .
6w 4C ns ns ** . . . .
8w 4C ns ns * ns . . .
10w 4C ns ns ns ns ns . .
12w 4C ns ns * ns ns ns .
14w 14C ns ns ns ns ns ns ns
- 13 -
Figures
Figure S1: Phylogenetic trees obtained from the analysis of DRM protein sequences. Physcomitrella
patens DNMT3 sequences were used as outgroup. Trees were obtained using maximum likelihood methods with 500 bootstrap replications and the best model to fit the alignment substitutions observed (evaluated using Mega6 software, version 6.06). The numbers in the nodes represent the percentage of trees where that branch is observed (bootstrap values), tree leafs are named by the protein name and in brackets the name of the species to whom the sequence belongs. In case of Physcomitrella patens sequences the leaves are highlighted with black dots before the leaf identifier. A: Tree obtained considering the complete protein sequences, using Jones-Taylor-Thornton model with a gamma distribution value of 1. B: Tree obtained from the sequences alignment and using Jones-Taylor-Thornton model with a gamma distribution value of 1.
Figure S2: Phylogenetic trees obtained from the analysis of DNMT3 protein sequences. Physcomitrella
patens DRM sequences were used as outgroup. Trees were obtained using maximum likelihood methods with 500 bootstrap replications and the best model to fit the alignment substitutions observed (evaluated using Mega6 software, version 6.06). The numbers in the nodes represent the percentage of trees where that branch is observed (bootstrap values), tree leafs are named by the protein name and in brackets the name of the species to whom the sequence belongs. In case of Physcomitrella patens sequences the leaves are highlighted with black dots before the leaf identifier. A: Tree obtained considering the complete protein sequences, using Jones-Taylor-Thornton model with a gamma distribution value of 4. B: Tree obtained from the sequences alignment and using the Whelan and Goldman model using a gamma distribution value of 3.
- 14 -
Figure S3: Picture of the 1 % agarose gel loaded with the individual PCR products for wild-type (WT)
and Δdrm2 mutant lines. The 1kb ladder (NEB) with the size of each of the ladder bands is presented on the sides of the image. A: samples from reactions A, using primers AT24 and AT26; B: samples from reactions B, using primers AT24 and AT25; C: samples from reactions C, with primers AT18 and AT27; D: samples from reactions D, with primers named CR7 and CR8. WT named samples were obtained from reactions using wild-type DNA as template from amplification; #1 samples were obtained from reactions using Δdrm2#1 line’s DNA, while #2 samples resulted from reactions where Δdrm2#2 line’s DNA was used as template. P named samples were obtained using pAT05 plasmid as template and B named samples used water instead of template DNA.
- 15 -
Figure S4: Average colony area of WT#1 colonies with 3, 5, 7 and 10 days of growth, germinated from
spores stored (0 to 14 weeks) at 4 ºC. Horizontal line represents samples’ average area and vertical bars represent standard error of the samples.
Figure S5: Average colony area of WT#1 colonies with 10, 15 and 21 days of growth, germinated from
spores stored (0 to 14 weeks) at 4 ºC. Horizontal line represents samples’ average area and vertical bars represent standard error of the samples.
Colony area of WT#1 (days 3-10)
WT#1 samples
Co
lon
y A
rea (
mm
2)
Fresh
d3
Fresh
d5
Fresh
d7
Fresh
d10
2w 4
C d
3
2w 4
C d
5
2w 4
C d
7
2w 4
C d
10
4w 4
C d
3
4w 4
C d
5
4w 4
C d
7
4w 4
C d
10
6w 4
C d
3
6w 4
C d
5
6w 4
C d
7
6w 4
C d
10
8w 4
C d
3
8w 4
C d
5
8w 4
C d
7
8w 4
C d
10
10w 4
C d
3
10w
4C d
5
10w
4C d
7
10w
4C d
10
12w 4
C d
3
12w 4
C d
5
12w
4C d
7
12w
4C d
10
14w
4C d
3
14w
4C d
5
14w 4
C d
7
14w 4
C d
10
0
1
2
3
4
5
6
Colony area of WT#1 (days 10-21)
WT#1 samples
Co
lon
y A
rea (
mm
2)
Fresh
d10
Fresh
d15
Fresh
d21
2w 4
C d
10
2w 4
C d
15
2w 4
C d
21
4w 4
C d
10
4w 4
C d
15
4w 4
C d
21
6w 4
C d
10
6w 4
C d
15
6w 4
C d
21
8w 4
C d
10
8w 4
C d
15
8w 4
C d
21
10w 4
C d
10
10w 4
C d
15
10w 4
C d
21
12w 4
C d
10
12w 4
C d
15
12w
4C d
21
14w
4C d
10
14w
4C d
15
14w
4C d
21
0
10
20
30
40
50
60
- 16 -
Figure S6: Average colony area of Δdrm2#1 colonies with 3, 5, 7 and 10 days of growth, germinated from
spores stored (0 to 14 weeks) at 4 ºC. Horizontal line represents samples’ average area and vertical bars represent standard error of the samples.
Figure S7: Average colony area of Δdrm2#1 colonies with 10, 15 and 21 days of growth, germinated from
spores stored (0 to 14 weeks) at 4 ºC. Horizontal line represents samples’ average area and vertical bars represent standard error of the samples.
Colony area of ∆drm2#1 (days 3-10)
∆drm2#1 samples
Co
lon
y A
rea (
mm
2)
Fresh
d3
Fresh
d5
Fresh
d7
Fresh
d10
2w 4
C d
3
2w 4
C d
5
2w 4
C d
7
2w 4
C d
10
4w 4
C d
3
4w 4
C d
5
4w 4
C d
7
4w 4
C d
10
6w 4
C d
3
6w 4
C d
5
6w 4
C d
7
6w 4
C d
10
8w 4
C d
3
8w 4
C d
5
8w 4
C d
7
8w 4
C d
10
10w
4C d
3
10w 4
C d
5
10w 4
C d
7
10w 4
C d
10
12w
4C d
3
12w
4C d
5
12w
4C d
7
12w
4C d
10
14w 4
C d
3
14w 4
C d
5
14w 4
C d
7
14w 4
C d
10
0
1
2
3
4
5
6
Colony area of ∆drm2#1 (days 10-21)
∆drm2#1 samples
Co
lon
y A
rea (
mm
2)
Fresh
d10
Fresh
d15
Fresh
d21
2w 4
C d
10
2w 4
C d
15
2w 4
C d
21
4w 4
C d
10
4w 4
C d
15
4w 4
C d
21
6w 4
C d
10
6w 4
C d
15
6w 4
C d
21
8w 4
C d
10
8w 4
C d
15
8w 4
C d
21
10w
4C d
10
10w
4C d
15
10w
4C d
21
12w 4
C d
10
12w 4
C d
15
12w 4
C d
21
14w 4
C d
10
14w 4
C d
15
14w 4
C d
21
0
10
20
30
40
50
60
- 17 -
Figure S8: Average colony area of WT#2 colonies with 3, 5, 7 and 10 days of growth, germinated from
spores stored (0 to 14 weeks) at 4 ºC. Horizontal line represents samples’ average area and vertical bars represent standard error of the samples.
Figure S9: Average colony area of WT#2 colonies with 10, 15 and 21 days of growth, germinated from
spores stored (0 to 14 weeks) at 4 ºC. Horizontal line represents samples’ average area and vertical bars represent standard error of the samples.
Colony area of WT#2 (days 3-10)
WT#2 samples
Co
lon
y A
rea (
mm
2)
Fresh
d3
Fresh
d5
Fresh
d7
Fresh
d10
2w 4
C d
3
2w 4
C d
5
2w 4
C d
7
2w 4
C d
10
4w 4
C d
3
4w 4
C d
5
4w 4
C d
7
4w 4
C d
10
6w 4
C d
3
6w 4
C d
5
6w 4
C d
7
6w 4
C d
10
8w 4
C d
3
8w 4
C d
5
8w 4
C d
7
8w 4
C d
10
10w 4
C d
3
10w 4
C d
5
10w 4
C d
7
10w 4
C d
10
12w 4
C d
3
12w 4
C d
5
12w 4
C d
7
12w 4
C d
10
14w 4
C d
3
14w 4
C d
5
14w 4
C d
7
14w 4
C d
10
0
1
2
3
4
5
6
Colony area of WT#2 (days 10-21)
WT#2 samples
Co
lon
y A
rea (
mm
2)
Fresh
d10
Fresh
d15
Fresh
d21
2w 4
C d
10
2w 4
C d
15
2w 4
C d
21
4w 4
C d
10
4w 4
C d
15
4w 4
C d
21
6w 4
C d
10
6w 4
C d
15
6w 4
C d
21
8w 4
C d
10
8w 4
C d
15
8w 4
C d
21
10w
4C d
10
10w 4
C d
15
10w 4
C d
21
12w 4
C d
10
12w
4C d
15
12w 4
C d
21
14w 4
C d
10
14w 4
C d
15
14w
4C d
21
0
10
20
30
40
50
60
- 18 -
Figure S10: Average colony area of Δdrm2#2 colonies with 3, 5, 7 and 10 days of growth, germinated
from spores stored (0 to 14 weeks) at 4 ºC. Horizontal line represents samples’ average area and vertical bars represent standard error of the samples.
Figure S11: Average colony area of Δdrm2#2 colonies with 10, 15 and 21 days of growth, germinated
from spores stored (0 to 14 weeks) at 4 ºC. Horizontal line represents samples’ average area and vertical bars represent standard error of the samples.
Colony area of ∆drm2#2 (days 3-10)
∆drm2#2 samples
Co
lon
y A
rea (
mm
2)
Fresh
d3
Fresh
d5
Fresh
d7
Fresh
d10
2w 4
C d
3
2w 4
C d
5
2w 4
C d
7
2w 4
C d
10
4w 4
C d
3
4w 4
C d
5
4w 4
C d
7
4w 4
C d
10
6w 4
C d
3
6w 4
C d
5
6w 4
C d
7
6w 4
C d
10
8w 4
C d
3
8w 4
C d
5
8w 4
C d
7
8w 4
C d
10
10w
4C d
3
10w 4
C d
5
10w
4C d
7
10w 4
C d
10
12w
4C d
3
12w
4C d
5
12w 4
C d
7
12w
4C d
10
14w
4C d
3
14w 4
C d
5
14w
4C d
7
14w
4C d
10
0
1
2
3
4
5
6
Colony area of ∆drm2#2 (days 10-21)
∆drm2#2 samples
Co
lon
y A
rea (
mm
2)
Fresh
d10
Fresh
d15
Fresh
d21
2w 4
C d
15
2w 4
C d
21
2w 4
C d
10
4w 4
C d
10
4w 4
C d
15
4w 4
C d
21
6w 4
C d
10
6w 4
C d
15
6w 4
C d
21
8w 4
C d
10
8w 4
C d
15
8w 4
C d
21
10w
4C d
10
10w
4C d
15
10w
4C d
21
12w
4C d
10
12w
4C d
15
12w
4C d
21
14w
4C d
10
14w
4C d
15
14w
4C d
21
0
10
20
30
40
50
60
- 19 -
Figure S12: 1 % agarose gel loaded with the in-tissue individual PCR reactions for genotyping of
selection-surviving colonies of transformation with pSP3b plasmid. 1kb ladder (NEB) was used to estimate the amplified fragments size (sides of the image). Wells numbered #1 to #10 represent the selection-surviving colonies number, the WT samples represents the wild-type (WT) tissue sample used as positive control for the reactions. P named wells were loaded with reactions using the plasmid DNA as template and B samples used DNA instead of template DNA. A: reactions using primers named SP31 and SP32; B: samples from reactions with primers SP32 and SP34; C: results from the reactions obtained using primers SP33 and SP35; D: samples from reactions using primers CR2 and CR3.