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Secondary Mutations Correct Fitness Defects in Toxoplasma gondii with Dinitroaniline

Resistance Mutations

Christopher Ma, Johnson Tran, Catherine Li, Lakshmi Ganesan, David Wood,

and Naomi Morrissette

Department of Molecular Biology and Biochemistry,

University of California, Irvine, Irvine CA, 92697

Genetics: Published Articles Ahead of Print, published on September 9, 2008 as 10.1534/genetics.108.092494

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Corresponding author:

Naomi Morrissette

Department of Molecular Biology and Biochemistry

University of California, Irvine

Irvine CA 92697-3900

(949) 824-9243 (phone)

(949) 824-8551 (fax)

nmorriss@uci.edu

Running title: Secondary Mutations in Tubulin

Key words: Apicomplexa, oryzalin, replication, microtubule

Abstract word count: 200

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ABSTRACT

Dinitroanilines (oryzalin, trifluralin, ethafluralin) disrupt microtubules in protozoa but not

vertebrate cells, causing selective death of intracellular Toxoplasma gondii parasites without

affecting host cells. Parasites containing α1-tubulin point mutations are dinitroaniline resistant

but show increased rates of aberrant replication relative to wild type parasites. T. gondii

parasites bearing the F52Y mutation were previously demonstrated to spontaneously acquire two

intragenic mutations that decrease both resistance levels and replication defects. Parasites

bearing the G142S mutation are largely dependent on oryzalin for viable growth in culture. We

isolated 46 T. gondii lines that have suppressed microtubule defects associated with the G142S

or the F52Y mutations by acquiring secondary mutations. These compensatory mutations were

α1-tubulin pseudorevertants or extragenic suppressors (the majority alter the β1-tubulin gene).

Many secondary mutations were located in tubulin domains that suggest that they function by

destabilizing microtubules. Most strikingly, we identified eight novel mutations that localize to

an eight amino acid insert that stabilizes the α1-tubulin M loop, including one (P364R) that acts

as a compensatory mutation in both F52Y and G142S lines. These lines have reduced

dinitroaniline resistance but most perform better than parental lines in competition assays,

indicating that there is a tradeoff between resistance and replication fitness.

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INTRODUCTION

Toxoplasma gondii is a human pathogen grouped within the Apicomplexa, a phylum of

protozoa which are all obligate intracellular parasites (LEVENE 1988). Infection with this parasite

can be life-threatening in immunocompromised individuals and cause birth defects or

miscarriage during fetal infection (BLACK and BOOTHROYD 2000). Other apicomplexans such as

Plasmodium spp. and Cryptosporidium spp. have considerable medical importance or are animal

pathogens that affect livestock. Apicomplexans have a specialized apex which contains unique

organelles that coordinate invasion of host cells (MORRISSETTE and SIBLEY 2002a). These

parasites are delimited by a pellicle, a composite structure formed by the association of the

plasma membrane with the inner membrane complex, an assemblage of flattened vesicles. The

invasive stages of apicomplexan parasites have two microtubule populations: subpellicular

microtubules and spindle microtubules. Subpellicular microtubules are non-dynamic; they

maintain both apical polarity and the characteristic crescent shape of the parasite by interacting

with the cytoplasmic face of the pellicle. Spindle microtubules form an intra-nuclear spindle to

coordinate chromosome segregation. Both populations are critically important to parasite

survival and replication.

Microtubules are built by the polymerization of α-β-tubulin heterodimers and typically

contain 13 protofilaments. Protofilaments are formed by longitudinal head-to-tail association of

heterodimers and laterally associate to assemble the microtubule lattice (DOWNING and NOGALES

1998). Biochemical and structural studies have elucidated the roles of tubulin domains which are

important to the regulation of microtubule polymerization and depolymerization. Both α- and β-

tubulins have H1-S2 (N) and M loops which coordinate contacts between adjacent

protofilaments (DOWNING and NOGALES 1998; LI et al. 2002; LOWE et al. 2001; NOGALES et al.

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1999; NOGALES et al. 1997). α-tubulin contains an eight amino acid insert missing from the

analogous region of β-tubulin which stabilizes the α-tubulin M-loop to promote protofilament

lateral association. Both α- and β-tubulins bind to GTP; however, only β-tubulin GTP can be

exchanged or hydrolyzed while α-tubulin GTP plays a structural role in this subunit (ANDERS

and BOTSTEIN 2001; DAVID-PFEUTY et al. 1977; NOGALES et al. 1998; SAGE et al. 1995a; SAGE

et al. 1995b). The β-tubulin subunits are stimulated to hydrolyze GTP in a polymerization-

dependent fashion by association of a GTPase activating domain from the α-tubulin subunit of

the adjacent dimer within the protofilament. Structural evidence indicates that GTP-bound

dimers have a conformation that facilitates protofilament contacts by both α- and β-subunits.

After hydrolysis, GDP-tubulin dimers have a kinked conformation that is believed to weaken

lateral contacts to promote microtubule disassembly (NOGALES and WANG 2006a; NOGALES and

WANG 2006b; NOGALES et al. 2003; WANG et al. 2005; WANG and NOGALES 2005).

Protozoan parasites are sensitive to dinitroaniline compounds which disrupt microtubules

to inhibit replication (BHATTACHARYA et al. 2002; BOGITSH et al. 1999; MAKIOKA et al. 2000;

STOKKERMANS et al. 1996; TRAUB-CSEKO et al. 2001). Given that these compounds are inactive

against vertebrate or fungal tubulins, the mechanism of dinitroaniline action is of particular

interest for development of new anti-microbial therapies. Our assays, as well as data from others,

indicate that wild type T. gondii parasites have an IC50 value of 0.25 μM for the dinitroaniline

oryzalin and display aberrant phenotypes in culture at 0.5 μM oryzalin (MORRISSETTE and

SIBLEY 2002b; STOKKERMANS et al. 1996). Although extracellular parasites are refractory to the

effects of microtubule-disrupting drugs, during intracellular growth, parasite microtubules are

dynamic and are sensitive to disruption by submicromolar concentrations of oryzalin or other

dinitroanilines.

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Dinitroaniline selectivity is associated with restricted binding to sensitive (plants and

protozoa) but not resistant (vertebrates and fungi) tubulin (CHAN and FONG 1990; HESS and

BAYER 1977; HUGDAHL and MOREJOHN 1993; MOREJOHN et al. 1987). Studies using molecular

dynamics analysis and compound docking indicate that dinitroanilines interact with a consistent

site on multiple conformations of α-tubulin derived from independent molecular dynamics

models of T. gondii, Leishmania spp. and Plasmodium spp. tubulins (MITRA and SEPT 2006;

MORRISSETTE et al. 2004). Parallel analysis of vertebrate (bovine) α-tubulin reveals that

dinitroanilines have non-specific, low affinity interactions and no consensus binding site,

consistent with in vivo and in vitro observations that these compounds do not bind to vertebrate

tubulin or disrupt vertebrate microtubules. This work predicts a binding site on parasite α-

tubulin that is located beneath the H1-S2 (N) loop. Since this loop participates in protofilament

interactions, the binding site location suggests that dinitroanilines disrupt lateral contacts in the

microtubule lattice. Consistent with this notion, a comparison of α-tubulin molecular dynamics

simulations in the presence and absence of bound drug suggests that dinitroaniline binding

profoundly limits flexibility of the α-tubulin H1-S2 loop which is drawn in towards the core of

the tubulin dimer (MITRA and SEPT 2006).

T. gondii is a haploid organism for most of its life cycle, although it is capable of sexual

recombination during a transient diploid zygote stage. There are three α- and three β-tubulin

genes in the genome: the α2- gene appears to be gamete stage specific and the divergent α3- gene

may be used to construct the conoid, an unusual structure built of tubulin sheets rather than

microtubules (HU et al. 2002). The α1- and β1-tubulin genes are the dominantly expressed

forms in the proliferative (tachyzoite) stage of the parasite studied here. We have previously

identified a number of α1-tubulin point mutations associated with oryzalin resistance in T. gondii

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(MA et al. 2007; MORRISSETTE et al. 2004). The mutations are distributed throughout the linear

sequence of α1-tubulin. However, when mapped onto a model of T. gondii α1-tubulin based on

the structure of the vertebrate tubulin dimer, many of the mutations cluster in specific domains of

the protein. For example, many mutations are located in or near domains that are associated with

microtubule stability or in the core of α-tubulin, adjacent to or within the dinitroaniline binding

site identified by computational methods. Our working model suggests that most tubulin

mutations confer dinitroaniline resistance by one of two possible mechanisms: (1) increasing

subunit affinity within the microtubule to compensate for the action of bound drug and (2) by

reducing the affinity of the tubulin for dinitroanilines by changes to the compound binding site.

Our previous observations that diverse mutations to α1-tubulin are sufficient to confer

resistance to dinitroanilines such as oryzalin in the protozoan parasite T. gondii might suggest

that dinitroaniline binding site ligands are not appropriate for development of anti-parasitic

agents. However, in many cases, drug resistance mutations isolated under laboratory conditions

are not observed in clinical settings since parasites are incapable of robust growth in natural

reservoirs. For example, a subset of laboratory selected dihydrofolate reductase (DHFR)

mutations that confer pyrimethamine resistance in P. falciparum are either rare or not observed

in the field (BATES et al. 2004; HANKINS et al. 2001; HASTINGS et al. 2002). Moreover, studies

that have exploited growth competition assays to assess the fitness of T. gondii strains bearing

wild-type or pyrimethamine resistant DHFR genes have concluded that even mutant strains that

behave similarly to wild-type parasites in vitro can display growth defects in vivo (FOHL and

ROOS 2003). We show here that dinitroaniline-resistant parasites have reduced fitness reflected

by increased rates of replication defects and poor performance in growth competition assays.

These lines rapidly and spontaneously acquire compensatory mutations that correct the fitness

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defects and reduce resistance to dinitroanilines. We conclude that drugs that selectively target

parasite microtubules remain a realistic option for the development of new anti-parasitic

therapies.

MATERIALS AND METHODS

Culture of Toxoplasma lines: T. gondii tachyzoites (the standard RH strain and mutants

derived from the RH strain) were propagated in human foreskin fibroblast (HFF) cells in DMEM

with 10% FBS as previously described (ROOS et al. 1994). Oryzalin (Riedel-deHaen, Germany)

stock solutions were made up in DMSO. Lines bearing oryzalin-resistance mutations were

propagated in 0.5 µM oryzalin to suppress selection of secondary mutations.

Selection of suppressors: T. gondii lines bearing allelic replacements of the F52Y or

G142S mutations to the α1-tubulin gene were generated as previously described (MA et al. 2007;

MORRISSETTE et al. 2004). These lines were serially passed for 3-4 weeks in the absence of

oryzalin to select for spontaneous mutants with improved growth. After single-cell cloning, the

clones displayed robust growth and each of the lines analyzed represents an independently

derived (non-sibling) lineage. Clonal lines were analyzed for changes to the α1- or β1-tubulin

genes.

Analysis of tubulin point mutations: The α1- and β1-tubulin genes were amplified from

genomic DNA isolated from individual parasite lines as previously described (MA et al. 2007;

MORRISSETTE et al. 2004). The α1-tubulin gene was amplified using thermal cycling with

primers GAGTCTCGTAGAGAACAAGC (5’ UTRA) and CGTTTATACCTTCACCTTTTC (3’

UTRA), and the purified fragment was sequenced as previously described. The β1-tubulin gene

was amplified with primers GTGGTGTTGCGCCTTC (5’ UTRB) and CGAGTGTTTAG-

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GACAGTGAC (3’ UTRB) and was sequenced with the following primers: (BT3E1)

CATTCTCCGCGATTCTC; (BT5E2) GTCCGGGTGTTCCTAC; (BTF2a) CATCATGGAG-

ACTTTCTCC; (BTR2a) GGAGAAAGTCTCCATGATG; (BTF2b) CAAAGAACATGATG-

TGCG; (BTR2b) CGCACATCATGTTCTTTG; (BT3E2) CTCGTCCATACC TTCACC;

(BT5E3) GAGATGGCACATTTAGTGTG; (BT3E3) CTCCCTCTTCCTCTGC; and (BT5E4)

CCGAGTATCAGCAGTACC. DNA sequences were analyzed using Sequencher software

(GeneCodes) to identify point mutations in the coding sequence of the α1-tubulin or β1-tubulin

genes. We also checked lines for α2-, α3-, β2- and β3-tubulin mutations but did not identify any

changes to these genes.

Structural models of T. gondii tubulin: Point mutations were mapped onto models of

T. gondii tubulin using PyMol (DELANO 2002). The models (containing a rebuilt H1-S2 loop)

were previously generated for computational studies (MITRA and SEPT 2006; MORRISSETTE et al.

2004).

Immunofluorescence staining: Extracellular parasites were fixed, permeablized and

stained in suspension as previously described (MORRISSETTE and SIBLEY 2002b). The T. gondii

plasma membrane was labeled with DG52 antibody (BURG et al. 1988) and detected with an

Alexa 594 secondary antibody (Invitrogen). DNA was visualized with DAPI. Suspension

samples were allowed to settle onto 35 mm dishes with glass coverslip insets (MatTek). Images

were collected on a Zeiss Axioskop using the Axiovision camera and software. Images were

exported and manipulated in Photoshop 8.0.

Quantification of replication defects: T. gondii were passed into HFF cells without

oryzalin selection and allowed to grow until complete host cell lysis. Extracellular parasites

were viewed in suspension in MatTek dishes with a coverslip inset using a 63X phase-contrast

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lens on a Zeiss Axioskop microscope. Images were captured as tif files and scored by counting

as previously described (MA et al. 2007). In order not to under-represent aberrant forms, we

counted the number of apical regions to establish total parasites lost from the parasite population

through replication defects.

Competition assays: T25 flasks with confluent HFF cells were inoculated with a 1:1 ratio

of RH strain-derived lines described here and wild type (RH strain) parasites expressing

cytosolic GFP (1x107 parasites from each line) (KIM et al. 2001) . After host monolayer lysis, a

new T25 flask with confluent HFF cells was inoculated with 1.0 ml of the lysed culture and the

remaining material (5ml) was used for flow cytometry analysis. Extracellular parasites were

purified away from host cell debris by passing lysate through a 3 μm filter. After parasites were

collected by centrifugation (all spins performed at 6,000 RPM for 6 min), they were fixed in

1.0% formalin (Sigma) for 1 min at RT. After a PBS wash, parasites were blocked in 10% (w/v)

BSA/PBS (Fisher Scientific, 30 min, 4oC) and surface labeled with DG52 anti-P30/SAG1 mouse

antibody (30 min, 4oC, 1:1000 in 10% (w/v) BSA/PBS) and a PE-Cy5.5 conjugated goat anti-

mouse secondary antibody (Invitrogen, 30 min, 4oC, 1:1000 in 10% (w/v) BSA/PBS). The total

number of parasites (PE-Cy5.5 labeling) and the number of parasites with GFP fluorescence

were enumerated by flow cytometry using a FACSCalibur flow cytometer (Becton Dickinson,

Inc.) and analyzed with Flowjo software (Tree Star, Inc.). The trend lines represent the

percentage of total mutant parasites at each time point, and the analysis was carried out for 7 and

15 serial parasite passages.

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RESULTS

Toxoplasma α1-tubulin mutations are associated with oryzalin resistance but have

fitness defects: We previously isolated a number of T. gondii lines that have mutations to α1-

tubulin which are sufficient to confer resistance to dinitroanilines such as oryzalin (MA et al.

2007; MORRISSETTE et al. 2004). Among the lines isolated in our work, are two that are oryzalin

resistant due to point mutations F52Y or G142S in the α1-tubulin gene (Figure 1). Parasites

bearing the G142S mutation are resistant to 1.0 μM oryzalin. G142 participates in binding the α-

phosphate portion of GTP (GIGANT et al. 2000; LOWE et al. 2001), and parasites with the G142S

mutation are largely dependent on the presence of oryzalin for viable growth in culture. F52 is

located in the H1-S2 loop, a domain that is critically important for protofilament-protofilament

associations in the microtubule lattice (LI et al. 2002). Parasites bearing the F52Y mutation have

high rates of defective replication and have been shown to spontaneously acquire compensatory

mutations. We previously reported that parasites with the F52Y mutation were resistant to 7 μM

oryzalin (MA et al. 2007). In the course of carrying out the experiments reported here, we

discovered that parasites bearing this mutation rapidly acquire secondary mutations (even when

under 0.5 μM oryzalin selection) and that the true resistance of the parental line is 13 μM.

Relative to wild type T. gondii, both F52Y and G142S parasite lines have high rates of overt

replication defects (Figure 1B and C).

Defects are corrected by compensatory mutations in lines derived from the two

parental parasite strains: When T. gondii parasites bearing an allelic integration of the F52Y or

G142S mutation are passed for several generations in the absence of oryzalin selection, they

spontaneously acquire mutations that allow them to grow with increased robustness. After

single-cell cloning independently selected lines with improved growth, we analyzed the

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sequences of T. gondii α1- and β1-tubulin genes for mutations associated with suppression of

defects. We identified both α1-tubulin pseudorevertants and extragenic mutations in β1-tubulin

and other (unidentified) genes. We isolated twenty-seven independent lines from the F52Y line

and identified mutations that localize to α1-tubulin or β1-tubulin in twenty-two of these lines

(Figure 2 and Table 1). Three of the α1-tubulin mutations are located in the M or H1-S2 (N)

loops and seven mutations are located in the eight amino acid α-tubulin specific insert. Six of

the lines have mutations to the β1-tubulin gene; three of these are in the M or H1-S2 (N) loops.

The G142S resistance mutation alters the GGGTGSG tubulin motif which is associated with

binding the phosphate portion of GTP. Of the twenty-four G142S derived lines characterized

here, thirteen acquire a secondary mutation in the α1-tubulin gene (Figure 3 and Table 2). One of

these mutations is associated with GTP binding (S171A), two localize in the eight amino acid α-

tubulin specific insert and one is in the M loop. We also isolated ten lines with extragenic

suppressors, with eight of these occurring in the β1-tubulin gene and two being in other,

unidentified loci. During the course of these studies we did not identify any true revertants of

F52Y or G142S parasites.

Growth competition assays indicate that the derived lines have increased fitness

relative to the parental strains: We have exploited an established GFP-expressing T. gondii

line to assess the relative fitness of the F52Y and G142S parasite lines and the derived progeny

(KIM et al. 2001). Both the GFP-expressing line and the tubulin mutant lines are derived from

RH strain T. gondii. We co-infected HFF cells with an equivalent number of GFP-expressing

parasites and parasites of each tubulin mutant line, and exploited green fluorescence to follow

the relative rates of replication and growth over serial passage (Figure 4). Parasites from lysed

flasks were used to inoculate new flasks such that they would lyse the host cell monolayer in 2

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days time. The remainder of each sample was analyzed by flow cytometry. In initial

experiments, the parasites were labeled with the DG52 (anti-SAG1) antibody to ensure that the

entire parasite population was visualized for analysis (Figure 4 inset). Once we were confident

that our gating captured the parasite population, we omitted the DG52 labeling, with consistent

results. At each time point, the total number of parasites and the number of GFP labeled parasite

were determined. Non-fluorescing parasites represent the fraction of the population expressing

the specific tubulin mutation(s). These growth competition assays indicate that the F52Y and

G142S lines compete poorly with wild-type-GFP parasites (Figure 4). After 6 days, the G142S

line is not detectable and at 8 days the F52Y line cannot be detected. In contrast, the β1-P61A

and α1-P364R lines which are derived from the G142S and F52Y parasites are still present at

this time, although their declining percentage in the population indicates that they are less fit than

the GFP-RH line. As a control, we also carried out growth competition for wild type RH

parasites versus the GFP-expressing parasites. Over time, we reproducibly observed that the

GFP-expressing parasites were at a disadvantage relative to the wild type RH T. gondii.

Therefore, the growth competition using GFP-RH parasites underestimates the fitness defects

associated with the dinitroaniline resistance mutations and the associated suppressed lines. We

analyzed all of the F52Y and G142S derived lines after two weeks (14 days) in serial passage

with GFP-RH parasites to assess relative fitness relationships in competition with wild type

parasites (Figure 5 and 6). In most cases, lines with compensatory mutations do not fully recover

robust growth. The most effective F52Y derived lines with respect to fitness are point mutations

at T51A (the α-tubulin H1-S2 loop), P364A (the α-tubulin insert) and G34S (the β-tubulin H1-S2

loop), which are still all less than 50% of the culture after 2 weeks (Figure 5 and Table 3). The

most effective G142S derived lines are S171A (an α-tubulin GTP binding site residue), L230V

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(located in helix 7 of α-tubulin) and V363A (in the α-tubulin insert). The L230V line is present

at ~50% of the culture after two weeks (Figure 6 and Table 3). However, since the GFP-RH

parasites have a fitness defect relative to unlabeled RH strain wild type parasites, it is still likely

that the L230V line would compete unfavorably with wild type T. gondii.

The derived lines have diminished oryzalin resistance relative to the parental

strains: The F52Y and G142S parasite lines display 13 and 1.0 µM oryzalin resistance,

respectively. Lines derived from these parental strains display reduced oryzalin resistance

(Figure 5 and 6 and Table 3). With respect to the F52Y derived lines, the P364A line retains the

highest level of oryzalin resistance (7 μM) and competes relatively well with the GFP-RH

parasites. Since the G142S line parasites have 1 μM oryzalin resistance, the range of resistance

observed in the derived lines is substantially lower. A number of the lines (the Q256H, S287P,

V363A mutations in α-tubulin and the P358R and A393V mutations in β-tubulin) do not have

greater resistance to oryzalin than that observed for wild type parasites (<0.5 μM by

morphological criteria). Lines with secondary mutations at S171A and L230V retain the highest

levels of oryzalin resistance in the context of the greatest degree of overall fitness in the

competition assay. All 46 lines bearing compensatory mutations are associated with diminished

oryzalin resistance.

DISCUSSION

Previous genetic studies have identified secondary mutations that correct tubulin

associated defects. In studies of colchicine resistance, intragenic mutations correct defects

associated with the D45Y mutation to β-tubulin which confers colchicine resistance in CHO cells

(WANG et al. 2004). This substitution is located in the H1-S2 loop and since it increases

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microtubule stability, cells that express it are hypersensitive to Taxol. Lines that suppress the

Taxol hypersensitivity and associated temperature sensitive defects have point mutations at

V60A (the H1-S2 loop) or Q292H (near the M loop) of β-tubulin that alter microtubule stability.

When wild type CHO cells express a tagged β-tubulin construct bearing the D45Y mutation, the

microtubule array is strikingly denser relative to the array observed with expression of wild type

tubulin alone. Inclusion of secondary mutations in the β-tubulin constructs reduces the

microtubule array making it similar to that observed with expression of wild type tubulin. When

β-tubulin constructs with the V60A and Q292H secondary mutations alone are expressed, the

microtubule arrays are dramatically reduced relative to those observed with wild type tubulin

expression. These observations are strikingly similar to our results described here. The F52Y

mutation is predicted to increase microtubule stability to confer dinitroaniline resistance, but also

causes increased rates of replication defects. Secondary mutations are located in regions such as

the H1-S2 loop, the M loop and the α-tubulin insert domain which are predicted to decrease

microtubule stability. We attempted to express GFP-tagged vertebrate tubulin with analogous

mutations in COS cells to visualize differential assembly of these tubulins. Unfortunately,

expression appears to be somewhat toxic and the patterns were too variable to be used to assess

relative stability of tubulins with these mutations (JT and NM, data not shown).

Many of the secondary mutations are identical or quite similar to previously identified

mutations that decrease microtubule stability in Caenorhabditis elegans and Arabidopsis

thaliana. C. elegans sensory neuron function requires unusual 15-protofilament microtubules

(CHALFIE 1982; CHALFIE et al. 1986; CHALFIE and THOMSON 1979; CHALFIE and THOMSON

1982; SAVAGE et al. 1989; SAVAGE et al. 1994). These microtubules are built from dimers

encoded by distinct α- (mec-12+) and β- (mec-7+) tubulin genes (FUKUSHIGE et al. 1999; GU et

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al. 1996; SAVAGE et al. 1989; SAVAGE et al. 1994). A number of mec-7 and mec-12 alleles have

been identified in screens for sensory neuron defects. In addition, mutations in Arabidopsis that

convert linear growth of plant parts (such as roots and stems) to a twisted phenotype have altered

cortical microtubule arrays. This twisting morphology can be phenocopied by treatment of wild

type plants with the microtubule inhibitor propyzamide. Many of the twisting mutants have

mutations to α- and β-tubulin mutations that decrease microtubule stability and increase plant

sensitivity to microtubule disrupting drugs (ABE and HASHIMOTO 2005; ABE et al. 2004; ISHIDA

and HASHIMOTO 2007; ISHIDA et al. 2007a; ISHIDA et al. 2007b; NAKAJIMA et al. 2004;

NAKAJIMA et al. 2006; SHOJI et al. 2004; THITAMADEE et al. 2002). The β-tubulin mutation

G34S (a F52Y suppressor) was previously identified as a mild, recessive mec-7 allele (SAVAGE

et al. 1989). Other mec-7 alleles (P61L/S, G269D, R318G and A393T) alter equivalent residues

to different substitutions in T. gondii β-tubulin (the F52Y suppressor G269V and the G142S

suppressors P61A, R318C and A393V). The G142S secondary mutation I355E in α-tubulin is

adjacent to the location of the mec-12 allele G354E. Moreover, the A393 residue is also

identified as a threonine substitution causing twisting in Arabidopsis β-tubulin (A394T). The

F52Y compensatory mutations V181I and K280N in α-tubulin and H396Q in β-tubulin and the

G142S mutation P348L (α-tubulin) are near the twisting mutations S180P, A180T, A281T and

T349I (α-tubulin) and A394T (β-tubulin). Lastly, the G142S suppressor Q291E is adjacent to a

second glutamine at 292 that is a mec-7 allele (Q292P). This residue also influences microtubule

stability in the context of epothilone resistance (Q292G) and suppresses hyperstabilized

microtubules (Q292H) in D45Y colchicine-resistant cells (HE et al. 2001; WANG et al. 2004).

One of the novel findings presented in this work is the identification of a set of eight

mutations that localize to the α-tubulin specific insert (TVVPGGDL). The compensatory

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mutations in the F52Y line that localize to the insert are T361P, P364A, P364R, G366R, D367V

and L368F and the G142S line mutations are V363A and P364R. To our knowledge, the insert

substitutions represent the first description of mutations located in this highly conserved domain.

We hypothesize that these substitutions impair the ability of the insert to promote M loop lateral

contacts with the adjacent protofilament. Indeed, the D367V mutation eliminates a salt bridge

that occurs with R229 (MA et al. 2007). Strikingly, the P364R mutation functions as a

compensatory mutation in both lines, albeit more successfully in the F52Y line (compare

competition results in Figures 5 and 6 and Table 3). Although the G142S/P364R line does not

compete effectively enough with wild type parasites to remain in the culture at 14 days, it was

isolated as a line with improved growth over the G142S line and at some subtle level is likely to

improve fitness of the parental line. This is also likely to be the explanation for the F52Y

suppressor A295G in competition at 14 days (Table 3).

The studies presented here show a correlation between acquisition of increased fitness

and the appearance of mutations to the α1- or β1-tubulin genes. While the improved fitness

observed in parasite lines derived from the F52Y and G142S parental strains is likely to be due

to secondary tubulin mutations, other (non-tubulin) mutations may also contribute to the

observed phenotypes of decreased resistance and enhanced growth. Indeed, five of the lines

derived from the F52Y suppressor screen and one line derived from the G142S suppressor screen

have improved growth but do not have altered α1-, α3-, β1-, or β3- tubulin genes suggesting that

non-tubulin compensatory mutations do arise. Moreover, although we can and have created

allelic replacements for two α-tubulin pseudorevertants in wild type parasites (MA et al. 2007),

we cannot create allelic replacements for the β-tubulin suppressors since we cannot select for

these in the absence of dinitroaniline selection. When we analyzed the allelic replacements for

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the α-tubulin pseudorevertants (F52Y/A273V and F52Y/D367V) in the competition assay, they

performed differently from the equivalent lines with these spontaneous mutations. Surprisingly,

in both cases the transgene versions of the lines had greater fitness levels than the spontaneous

mutant lines (data not shown). The suppressed lines described here were isolated after 4-6 weeks

of growth in media without oryzalin. This represents the shortest time in which we could observe

parasites with new mutations. In contrast, it takes at least 6 weeks to isolate transgene-bearing

lines. We predict that further growth of lines with impaired fitness in the absence of oryzalin

selection would select for additional mutations that collectively contribute to increased fitness.

Previous studies have established that there is a tradeoff between carrying genes that

confer drug resistance and the fitness cost of these altered alleles, particularly in the absence of

drug selection. Recent field studies from Malawi indicate that Plasmodium falciparum parasites

in this region lost resistance to chloroquine in fewer than ten years after sulfadoxine-

pyrimethamine treatment replaced chloroquine treatment for individuals suffering from malaria

(LAUFER et al. 2006; VOGEL 2006). These field observations, as well as the controlled laboratory

studies of DHFR mutations in P. falciparum and T. gondii described in the Introduction, indicate

that the existence of mutant lines that express resistant forms of target proteins does not rule out

the development and use of agents that inhibit these proteins. With respect to the studies

presented here, we believe that although dinitroanilines or other microtubule disrupting

compounds do not yet exist in an appropriate form for treatment of parasite infections,

selectively targeting protozoan tubulin remains a viable strategy for eliminating parasite

infections.

19

ACKNOWLEDGEMENTS

We thank Dan Sackett for commenting on this manuscript, David Sibley and Michael

Grigg for helpful conversations on competition assays and Sheryl Tsai bringing clarity to

PyMOL commands. Research presented in this paper was supported by NIH grants AI055790

and AI067981 to NSM. LG and DW are undergraduate students in the campus-wide honors

program at UCI and LG was supported by a UCI SURP fellowship. We would like to thank

Candice Kwark and Brian Luk, high school student participants in the American Cancer Society

summer research program at UCI, who helped select and analyze the lines described here.

20

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FIGURE LEGENDS

FIGURE 1. — (A) The F52Y and G142S α-tubulin point mutations confer 13 and 1 µM oryzalin

resistance when integrated into wild type (sensitive) T. gondii parasites. The F52Y mutation

(red) is located in the H1-S2 (N) loop (red) of α1-tubulin. The computationally determined

dinitroaniline binding site is below this loop (oryzalin shown in orange). Interactions between

the H1-S2 (N) loop and the M loop of adjacent dimers coordinate lateral associations between

protofilaments in the microtubule lattice. The G142S mutation is located in the core of α1-

tubulin and the G142 residue represents the initial glycine in the GGGTGSG motif (blue) which

is characteristic of tubulins. This motif is part of the GTP binding site which is essential for

correct folding of the tubulin dimer and interacts with the phosphate portion of GTP (purple). (B)

Quantification of replication defects in the absence of oryzalin. The results for F52Y parasites

were previously published in (MA et al. 2007) and are shown here for purposes of comparison

with the G142S line. (C) Immunofluorescence images of extracellular parasites of the G142S

line labeled with DG52 (red, labels the parasite surface) and DAPI (blue, labels the parasite

DNA). The lower panel shows DNA distribution only. (1) A normal appearing parasite has a

crescent shape and DNA staining of the nucleus (arrow) and apicoplast DNA (arrowhead). (2-6)

Parasite “monsters” result from replication defects due to improper coordination of spindle

microtubule (nuclear division) and subpellicular microtubule (cytokinesis) functions. Although

the extracellular parasites in panels 3 and 4 are similar to intracellular replicating stages (see

diagram in (MA et al. 2007)), parasites can not complete division outside of host cells and their

abnormal shape prevents invasion of new host cells. In addition, panels 2, 5 and 6 illustrate

parasites with diffuse (2), improperly segregated (3 nuclei, panel 5) and non-segregated (6)

nuclear staining (compare to panel 1).

25

FIGURE 2. — Location of compensatory mutations obtained from the F52Y line mapped on a

model of the Toxoplasma tubulin dimer. The α1-subunit is colored white and the β1-subunit is

blue. (A) Most mutations fall between the M and H1-S2 loops (red) facing the inner lumen of

the microtubule. The F52Y mutation is located in the H1-S2 loop of α1-tubulin (red) and

secondary mutations in α1-tubulin (black text) occur at T51A, V181I, A273V, K280N, N293D,

A295G, S300T, M313T, P360A, T361P, P364A, P364R, G366R, D367V, L368F and R373C

(orange/yellow). Mutations in yellow localize to the α-tubulin specific insert. Extragenic

suppressors in the β1-tubulin subunit (white text) occur at G34S, G82D, P261S, G269V, L273V

and H396Q. (B) The β1-tubulin mutation P261S is the only mutation that faces the outer surface

of the dimer within the microtubule lattice, while the β-subunit mutation H396Q is located at the

dimer-dimer interface within the protofilament. (C) The mutations T361P, P364A, P364R,

G366R, D367V and L368F occur in the α-tubulin specific insert (yellow) and the mutations

P360A and R373C are adjacent to this insert (orange).

FIGURE 3. — Location of compensatory mutations obtained from the G142S line mapped on a

model of the Toxoplasma tubulin dimer (the α1-subunit is white and the β1-subunit is blue). The

G142S mutation (yellow) is located in the GTP binding domain of α-tubulin. (A) Most

mutations (orange/yellow) are on the surface of the dimer that faces the inner lumen of the

microtubule. Secondary mutations in α1-tubulin (black text) occur at I93V, G131R, S171A,

I209V, L230V, Q256H, S287P, P348L, I355E, V363A, P364R, V371G and F418I. Suppressors

in the β1-tubulin subunit (white text) occur at residues A18P, P61A, F85L, K122E, H264Q,

Q291E, R318C, N337T, P358R, M363V and A393V. G142S α1-tubulin mutations at V363A

26

and P364R occur in the α-tubulin specific insert (yellow). (B) The α1-tubulin mutation at Q256

and the β1-tubulin mutation at H264 are the only mutations that face the outer surface of the

dimer within the microtubule lattice. The β-subunit mutation A393V is located at the dimer-

dimer interface within the protofilament. (C) The primary resistance mutation at G142 is located

within the tubulin motif (teal) of the α-tubulin GTP binding site. The GTP moiety is colored

blue and residues contributing to GTP binding outside of the GGGTGS tubulin motif are colored

red. The primary mutation G142S is located in the binding site and interacts with the α-

phosphate. The S171A mutation also locates to the binding site; S171 interacts with the ribose

portion of GTP (GIGANT et al. 2000; LOWE et al. 2001).

FIGURE 4. — In order to assess relative fitness of the F52Y and G142S lines as well as the

strains derived from these lines, equivalent numbers of parasites from a specific mutant line and

wild type RH strain parasites expressing GFP were inoculated into culture and the relative

percentage of the each population was quantified over time during serial passage using flow

cytometry. The relative parasite concentrations were quantified at the time of complete host cell

lysis when the lysate was passed into a new flask of host cells containing parasites. The red line

indicates the behavior of G142S parasites in competition and the orange line is the G142S

suppressor line β-P61A (located in the β-tubulin H1-S2 loop). The green line follows a

competition assay containing F52Y and the teal line represents behavior of the F52Y secondary

mutation α-P364R (located in the α-tubulin specific insert). The G142S parasites are out-

competed by GFP-RH parasites by day 6 whereas the β-P61A suppressor line persists until day

24. The F52Y parasites are eliminated at day 12 but in the presence of the α-P364R secondary

mutation, they are still present, albeit at reduced levels, when the competition was terminated at

27

30 days. The purple line traces the control competition between wild-type (untransfected RH

strain parasites) and the GFP-transfected “wild-type” RH strain line. Unlabeled parasites have a

growth advantage over GFP-transfected parasites. (Inset) A representative FACS dot plot of

DG52-labeled (red) parasites showing relative numbers of GFP-expressing wild type

Toxoplasma and a mutant line after a single passage.

FIGURE 5. — Replication fitness and oryzalin resistance in F52Y parasites and the derived

lines. Black bars: the percentage of mutant parasites relative to GFP-RH parasites after 7

passages (14 days). The assays were carried out in triplicate and error bars indicate standard

error of the mean. Gray bars: oryzalin resistance levels observed for F52Y parasites and the

derived lines.

FIGURE 6. — Replication fitness and oryzalin resistance in G142S parasites and the 24 derived

lines. Black bars: the percentage of mutant parasites relative to GFP-RH parasites after 7

passages (14 days). The assays were carried out in triplicate and error bars indicate standard

error of the mean. Gray bars: oryzalin resistance levels observed in G142S parasites and the

derived lines.