Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid...

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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

Transcript of Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid...

Page 1: Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid specific DNA methyltransferases in Physcomitrella patens ... debater e ter conversas parvas,

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

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“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

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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 -

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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

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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

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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

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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

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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

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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).

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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).

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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

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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).

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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).

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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

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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

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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

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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

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(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

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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.

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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

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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

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(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.

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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

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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).

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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

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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.

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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

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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.

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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

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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

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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

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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).

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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.

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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

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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.

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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.

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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

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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

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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).

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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

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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

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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

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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’

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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).

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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

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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

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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.

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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.

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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.

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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

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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

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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

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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

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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

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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

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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

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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.

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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

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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

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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

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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.

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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)

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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.

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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.

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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.

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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.

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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.

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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

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(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

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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

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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.

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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.

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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

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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.

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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.

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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

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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

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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

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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

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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 ***

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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

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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

Page 95: Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid specific DNA methyltransferases in Physcomitrella patens ... debater e ter conversas parvas,

- 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

Page 96: Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid specific DNA methyltransferases in Physcomitrella patens ... debater e ter conversas parvas,

- 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 ***

Page 97: Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid specific DNA methyltransferases in Physcomitrella patens ... debater e ter conversas parvas,

- 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

Page 98: Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid specific DNA methyltransferases in Physcomitrella patens ... debater e ter conversas parvas,

- 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

Page 99: Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid specific DNA methyltransferases in Physcomitrella patens ... debater e ter conversas parvas,

- 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

Page 100: Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid specific DNA methyltransferases in Physcomitrella patens ... debater e ter conversas parvas,

- 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.

Page 101: Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid specific DNA methyltransferases in Physcomitrella patens ... debater e ter conversas parvas,

- 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.

Page 102: Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid specific DNA methyltransferases in Physcomitrella patens ... debater e ter conversas parvas,

- 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

Page 103: Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid specific DNA methyltransferases in Physcomitrella patens ... debater e ter conversas parvas,

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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

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- 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

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- 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

Page 106: Deciphering the role of antherozoid specific DNA ... · Deciphering the role of antherozoid specific DNA methyltransferases in Physcomitrella patens ... debater e ter conversas parvas,

- 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.