Chromosomes of parasitic protist Trichomonas vaginalis and the

Chromosomes of parasitic protist Trichomonas vaginalis
Zuzana Zubáčová
Ph.D. thesis
Thesis supervisor: Prof. RNDr. Jan Tachezy, Ph.D.
Prague 2011
Department of Parasitology
Faculty of Science, Charles University
1
Výsledky prezentované v doktorské práci Mgr. Zuzany Zubáčové jsou společným dílem
pracovníků Laboratoře biochemické a molekulární parazitologie a spolupracujících tímů.
Prohlašuji, že autorka se na jejich získání zasloužila významným dílem.
Data presented in this thesis resulted from team collaboration at the Laboratory of
biochemical and molecular parasitology and during projects with our partners. I declare that
the author contributed substantially to obtaining of these results.
2
THERE´S A LOT GOIN´ ON…
Poďakovanie:
Ďakujem Honzovi Tachezymu za trpezlivé vedenie mojej práce a kontrolu textu dizertácie. Za
krásne roky spolupráce a pomoc ďakujem kolegom a kamarátom z laborky. Ďakujem rodičom
a Oliverovi za podporu v kritických chvíľach. Ďakujem Karlovi za poskytnuté zázemie
v závere práce.
3
Chromosomes of Trichomonas vaginalis
Contents
Abstract
6
1. Overview
7
1. 1. Introduction
7
1. 2. Nuclear genomes and chromosomes of parasitic protists
8
1. 2. 1. Genomes of Apicomplexa (CHROMALVEOLATA)
9
Plasmodium falciparum
9
Toxoplasma gondii
11
Cryptosporidium
11
1. 2. 2. Genomes of Microsporidia (OPISTHOKONTA)
12
1. 2. 3. Genome of Entamoeba histolytica (AMOEBOZOA)
13
1. 2. 4. Genomes of Kinetoplastida (EXCAVATA)
13
Trypanosoma brucei
13
Leishmania major
14
1. 2. 5. Genome of Giardia intestinalis (Metamonada, EXCAVATA)
15
1. 2. 6. Genomes of Trichomonas vaginalis and related species (Parabasala,
EXCAVATA)
16
2. Mitosis in parasitic protists
18
2. 1. Mitosis in Apicomplexa
20
2. 2. Mitosis in Kinetoplastida
22
2. 3. Mitosis in Microsporidia
24
2. 4. Mitosis in Entamoeba histolytica
24
2. 5. Mitosis in Giardia intestinalis
26
2. 6. Mitosis in Trichomonas vaginalis
27
References
29
3. Aims
39
4. Publications
40
 Draftgenome sequence of the sexually transmitted pathogen Trichomonas
vaginalis.
 Comparative analysis of trichomonad genome sizes and karyotypes.
 Microsatelite polymorphism in the sexually transmitted human pathogen
Trichomonas vaginalis indicates a genetically diverse parasite.
4
 Fluorescence in situ hybridization (FISH) mapping of single copy genes on
Trichomonas vaginalis chromosomes.
5. Unpublished data
41
 Histone H3 variants in Trichomonas vaginalis
6. Thesis summary
41
42
5
Abstract
Genome sequencing and associated proteome projects has revolutionized research in
the field of molecular parasitology. Progress in sequencing of parasite as well as free-living
species enables comparative and phylogenetic studies and provides important data sets for
understanding of basic biology and identification of new drug targets. The genome
sequencing of Trichomonas vaginalis revealed surprisingly large genome size of this
organism (~160 Mb) that resulted from expansion of various repetitive elements, specific
gene families and large scale genome duplication. I studied genome sizes of other nine
selected species from Trichomonadea group to find whether other trichomonads have
undergone similar genome expansion. The measurement of nuclear DNA content by flow
cytometry revealed relatively large DNA content in all tested species although within a broad
range (86-177 Mb). The largest genomes were observed in the Trichomonas and
Tritrichomonas genera while Tetratrichomonas contains the smallest genome. The genome
sizes correlated with the cell volume however no relationship between genome size and
pathogenicity, the site of the infection or trichomonad phagocytic ability was observed.
The published information about chromosomes of trichomonads was insufficient or
contradictory. To elucidate the number and appearance of chromosomes I investigated
karytypes of T. vaginalis and related trichomonads using conventional cytogenetic techniques
and fluorescence in situ hybridization (FISH). Karyotype analysis of selected trichomonad
species showed that all trichomonads have a single set of chromosomes that condense during
mitotic metaphase. The number of chromosomes in trichomonads ranges from four to six.
FISH analysis using probes against SSU and LSU rRNA genes showed that all 250 rDNA
units are localized to a single chromosome in one secondary constriction.
I also adapted FISH for T. vaginalis that is sensitive enough to detect single copy
genes on metaphase chromosomes. Sensitivity of conventional FISH was increased using
tyramide signal amplification. Our FISH protocol provides amenable tool for physical
mapping of the T. vaginalis genome and other essential applications such as development of
genetic markers for T. vaginalis genotyping with application in evolutionary studies,
comparative genomics and clinical praxis.
Similar to the other eukaryotes, successful mitosis is essential for reproduction of T.
vaginalis, however many aspects of this process are not fully understood. In my thesis, I
summarized current understanding of mitosis of T. vaginalis including my preliminary results
of study of trichomonad centromeres. I studied chromatin localization of T. vaginalis histone
H3 variants by transfecting trichomonads with their tagged versions. Among 23 of T.
vaginalis histones H3, I identified the homolog of cenH3 variant, which is the universal
marker of eukaryotic centromeres.
6
1. Overview
1.1. Introduction
With the beginning of the 21st century started the genomic era by deciphering of
human genome (Venter, 2001). It opened thousands new doors for basic as well as clinical
research. Knowledge of whole genomes fundamentally improved our understanding of
biology of pathogenic organisms including parasitic protist Trichomonas vaginalis. T.
vaginalis is a parasitic member of Parabasala group that belongs to eukaryotic supergroup
Excavata. It is an anaerobic organism, which invades the urogenital tract and causes the most
frequent non-viral sexual transmitted disease in humans called trichomoniasis. Despite its
medical importance, T. vaginalis was for a long time rather understudied organism (Carlton et
al. 2007). Its popularity as a subject of basic research started to grow after the release of data
from genome sequencing project in 2007. The summary of the results is published in Carlton
et al. (Carlton et al. 2007). Analyses of sequence data and subsequent proteomic studies
revealed new biochemical pathways in trichomonads, provided information for development
of new genetic markers for population studies that have potential to explain the mechanism of
drug resistance, allowed identification of new virulence markers and many more. The most
surprising result of Trichomonas genome project was the genome size. Although the predicted
genome size was about 25 Mb (Wang and Wang 1985), the genome sequence analysis
revealed the size of about 160Mbp coding for about 60 000 genes. Thus, Trichomonas
vaginalis appeared to be a parasitic protist with the largest genome and higher gene coding
capacity than human. Observed large genome size resulted from large scale genome
duplication events, accumulation of various repetitive elements and expansion of specific
gene families (Carlton et al. 2007).
T. vaginalis is a haploid, clonal protist that reproduces by mitosis. Ultrastructural
studies revealed that during mitosis of T. vaginalisthe nuclear envelope remains intact. The
mitotic spindle is formed extranuclearly with kinetochoral microtubules that interact with
kinetochores of chromosomes via nuclear envelope barrier (Brugerolle, 1975). This mitosis
variant is named cryptopleuromitosois (Raikov, 1994) and it was observed exclusivelly in
parabasalids. Although meiotic division has not been described in Trichomonas, figures of
chromosomes typical for meiotic division were rarely observed on chromosomal preparations
by Drmota and Kral (Drmota and Kral, 1997). In addition, eight of nine meiosis specific
genes were found in its genome suggesting that T. vaginalis is eighter currently sexual or just
recently asexual organism (Malik et al., 2008). During mitosis, T. vaginalis genome and also
genomes of other trichomonads condense into tiny well-structured haploid chromosomes. The
7
chromosome number is stable within trichomonad species and vary from four to six between
tested species (Zubacova et al., 2008). According to Drmota and Kral (Drmota and Kral,
1997), who described constrictions on T. vaginalis chromosomes as centromeres, T. vaginalis
has monocentric rather than holocentric chromosomes. However, mechanism of chromosome
segregation into next cell generation has not been studied in T. vaginalis. The structure of T.
vaginalis kinetochores, the mode of interaction between kinetochores and mitotic
microtubules, and the role of nuclear envelope in segregation of chromosomes is still unclear.
1. 2. Nuclear genomes and chromosomes of parasitic protists
A term genome means total hereditary information of an organism. Nuclear genomes
are compacted into cell structures called chromosomes. Chromosomes consist of DNA
associated with proteins, first of all histones, that enable folding and packing of long DNA
molecule into nucleus. Complex of DNA and associated proteins is called chromatin (the
Greek chroma means color and reflects staining properties of chromatin). In chromatin, DNA
is wound around complex of core histones consisting of histones H2A, H2B, H3 and H4 and
its specialized variants. Histone H1 binds linker DNA between nucleosomes. Histones were
most likely inherited from archaebacterial ancestors at early stage of eukaryotic evolution
(Talbert and Henikoff, 2010). Although histones belong among the most conserved proteins,
histones of parasitic protists show remarkable divergence from homologs of higher eukaryotes
what rise the question how these differences influence chromatin structure and condensation.
Significant sequence deviations were observed for example in Trypanosoma brucei (Bender
et al, 1992), Trypanosoma cruzi (Bontempi et al., 1994), Entamoeba histolytica (Födinger et
al., 1993), Entamoeba invadens (Sanchez et al., 1994), Leishmania infantum (Soto et al.,
1994), T. vaginalis (Marinets et al, 1996) and Giardia intestinalis (Wu et al. 2000).
During the cell cycle chromosomes usually pass through cycles of condensation
(strongly packed and displaying distinct morphology) and decondensation (chromatin is
relaxed in the nucleus with no obvious morphology). A species or an individual specific set of
chromosomes with characteristic number and appearance is called karyotype. In parasitic
protists number of chromosomes varies from five in Tritrichomonas foetus to hundred of
minichromosomes in Trypanosoma brucei (Alberts et al., 2002; Drmota and Kral, 1997;
Ersfeld, 2011). However, in some parasites such as trypanosomatids (Gull et al., 1998; Ravel
et al., 1998; Dubessay et al., 2004) and Entamoeba histolytica (Willhoeft and Tannich, 1999),
chromosomes do not condense in any state of their life cycle. Chromosomes can not be
visualised using standard cytogenetic staining also in Plasmodium (Coppel and Black, 2005).
8
Number of genome copies that is usually related to number of chromosomes in an organism
determines its ploidy. Parasitic protists are often haploid not only regarding genome copies
but also chromosome segments. For example, majority of Trypanosoma brucei genome is
diploid but regions of the genome containing VSG genes are haploid. These regions lacks
homologous chromosomal segments (Melville et al., 2000; Nuismer and Otto, 2004 ). The
evolutionary advantage of haploidy in parasites has been proposed in regard to successful host
infection as alleles connected with successful invasion of a host tend to be recessive. Haploid
group of parasites includes important pathogens of plants such as Plasmodiophora brassicae
and Spongospora subterranean and human protist pathogens such as Plasmodium falciparum,
Toxoplasma gondii and Trichomonas vaginalis (Nuismer and Otto, 2004).
Data from parasite´s genome projects present enormous amount of information in
regard to basic research of their biology and uncovering of new strategies for development of
antiparasital drugs (Coppel and Black, 2005). Access to genomic datasets of individual
genome projects of pathogenic protists is currently enabled via public databases including
National Center for Biotechnology Information (NCBI).Genomes of human pathogens are
integrated into database EuPathDB (ttp://eupathdb.org/eupathdb/, (Aurrecoechea et al., 2005))
with elaborated system of bioinformatic tools. This database serves as an entry point for
study of individual genes and genomes of Plasmodium (PlasmoDB), Toxoplasma (ToxoDB),
Cryptosporidium (CryptoDB), Trypanosoma and Leishmania (TriTrypDB), Entamoeba
(AmoebaDB),
Encephalitozoon
and
Enterocytozoon
(MicrosporidiaDB),
Giardia
(GiardiaDB) and Trichomonas (TrichDB).
1.2.1. Genomes of Apicomplexa (CHROMALVEOLATA)
Plasmodium falciparum
P. falciparum causes deadly human disease malaria that in 2009 took 781 000 lives. In
Africa,
one
in
every
five
children
deaths
is
due
to
malaria
(http://www.who.int/topics/malaria/en/). The genome project had the major ambition to
accelerate the research in direction of new strategies for elimination of malaria using new
chemotherapeutic or vaccination.
The size of nuclear genome of malaria parasite is ~23 Mb (Gardner et al., 2002). It consists
of haploid set of 14 chromosomes that do not condense. Conventional cytogenetic spreading
techniques for chromosomes imaging do not lead to visualization of Plasmodium
9
chromosomes (Gerald et al, 2011; Kelly et al., 2006; Prensier and Slomianny, 1986). The
chromosomes have diffuse appearance when stained for fluorescence microscopy and it is
difficult to observe these chromosomes using transmission electron microscopy. However,
chromosomes of P. falciparum can be resolved by pulse field gel electrophoresis (PFGE).
This technique is widely used in parasites
for determining the number and size of
chromosomes (electrophoretic karyotype) (Coppel and Black, 2005; Kelly et al. 2006;
Prensier and Slomianny 1986). P. falciparum chromosomes considerably vary in their length
due to variations in subtelomeric regions. Natural variation in chromosome length between
Plasmodium isolates results from meiotic recombination of parasites in the mosquito vector.
Chromosome size variation can be observed also in laboratory cultures. However, it this case
the variation is caused by chromosome breakage and reparation events that are independent
on meiotic recombination (Coppel and Black, 2005;Gardner et al. 2002).
Genome of P. falciparum is the most AT – rich genome sequenced to date. The
overall A/T content is 80, 6%. The most AT-rich regions are present in introns, intergenic
regions and at the site of putative centromeres. In the genome sequence, 5286 protein coding
genes were predicted, from which 54% contain intrones. No transposones or
retrotransposones were detected (Gardner et al., 2002).
Plasmodium genome is characteristic with unusual organization of rDNA units.
Ribosomal RNA coding genes and their arrangement are generally one of the most
extensively studied part of the genome (Prokopowich et al. 2003). In most eukaryotes rDNA
units are organized as a tandemly repeated arrays of 18S – 5,8S – 28S rRNA genes that are
clustered in nucleolar organizer region (NOR) on one or more chromosomes. The numbert of
rDNA copies usually positively correlates with the genome sizes of eukaryotic genome
(Prokopowich et al. 2003). However, units of ribosomal RNA genes in Plasmodium are not
clustered but dispersed throughout different chromosomes and their copy number is
unusually low: 4-8 per genome (Waters, 1994). Furthermore, Plasmodium rDNA units are
not identical in their sequences and their expression is regulated developmentally when
different set of rRNA is expressed at different stages of the parasite life cycle. It has been
proposed that Plasmodium is able to alter the rate of the gene expression by changing the
properties of ribosomes resulting in change of cell development (Gardner et al. 2002).
Genes responsible for virulence, adhesion molecules and antigenic variation are
present in multiple copies and are typically localized in subtelomeric regions of
chromosomes. Deletions in subtelomeric regions alter expression of adhesive molecules
10
leading to the lost of cytoadherence of infected erythrocytes to endhotelial cells (Day et al.
1993; Gardner et al. 2002).
Toxoplasma gondii
T. gondii is an important human and veterinary pathogen that causes toxoplasmosis. It
is estimated that about one-third of the human population is chronically infected that make T.
gondii one of the most prevalent parasites in humans (Laliberté and Carruthers, 2008). Most
infections of T. gondii are asymptomatic. However, it may propagate in immunocompromised
patiens such as individuals with Acquired Immune Deficiency Syndrome that may lead to
damage of various organs and even death. In pregnant women, T. gondii could be
transplacentally transmitted to the foetus that results in development of serious defects
(Dubey et al., 1998). Toxoplasma can be easily propagated in laboratory in human foreskin
fibroblasts (Roos et al., 1994) and methods for reverse genetics are available (Brooks et al.
2011). Thus Toxoplasma has became a model apicomplexan organism. The B7 strain that is
commonly associated with AIDS patiens was used for analysis of Toxoplasma genome. The
genome size of T. gondii is 80 Mb and contains 8000 protein coding genes (Coppel and
Black, 2005; Kissinger et al., 2003; Xia et al., 2008). Initial studies of karyotype using PFGE
revealed 11 chromosomes in Toxoplasma (Kissinger et al. 2003; Sibley and Boothroyd,
1992), however, more recent observation showed that
T. gondii genome consist of 14
chromosomes (Coppel and Black, 2005). In comparison to genes of Plasmodium or
Cryptosporidium, T. gondii genes are richer in introns. Genes coding secretion proteins of
apical complex organelles are located typically in subtelomeric regions of chromosomes
(Coppel and Black, 2005). rRNA genes that comprise of 110 units per haploid genome are
organized conventionally as tandem repeats (Guay et al. 1992).
Cryptosporidium
C. parvum and C. hominis are two cryptosporidial species that cause infection of
humans. They differ in the host range, genotype and pathogenicity. C. hominis is a human
specific parasite, while C. parvum infects also other mammals (Xu et al., 2004).
Cryptosporidiosis belongs to a group of opportunistic infections pathogens. In
immunocompromised patients it may cause serious chronic diarrhea resulting even in death
(O´Connor et al., 2011). No long-term in vitro cultures of Crysptosporidium are available.
11
However, large amount of Cryptosporidium cysts for experiments can be obtained from the
feces of infected calves and then be in vitro excysted (LaGier et al., 2003).
The genomes of both human species C. parvum and C. hominis were sequenced
however only subtle differences were found. The 9 Mb genome of Cryptosporidium consists
of eight chromosomes that were examined by molecular karyotyping using PFGE
(Abrahamsen et al., 2004; Blunt et al., 1997; Xu et al., 2004). Similarly to microsporidia,
genomes of Cryptosporidium are highly compact , however forces that affected their genome
sizes were different (Keeling, 2004). There are two basic mechanisms leading to genome
reduction: (i) shortening of intergenic regions, and (ii) elimination of genes from the genome
(Keeling, 2004). Genome of Cryptosporidium was affected predominantly by the first
mechanism. It is characteristic by reduction of intergenic regions and introns as well as
genes are reduced in their length. In the genomes of C. parvum and C. hominis, 3807 and
3994 genes were predicted, respectively, that is less than in Plasmodium or Toxoplasma. In
contrast to other apicomplexan, subtelomeric gene clusters encoding variant surface proteins
were not detected in Cryptosporidium (Coppel and Black, 2005).
Genes coding rRNA are organized according to the plasmodial paradigm. They are
present in five copies and they are not clustered in the genome. The rDNA units are dispersed
throughout the genome at least on three chromosomes. Like in Plasmodium, rDNA units
accumulated sequence heterogeneities propably because of dispersion in the genome (Le
Blancq et al., 1997).
1.2.2. Genomes of Microsporidia (OPISTHOKONTA)
Genomes of microsporidia range from 19,5 Mb in Glugea atherinae to 2.9 Mb in
Encephalitozoon cuniculi and 2,3 Mb in Encephalitozoon intestinalis that is the smallest
known eukaryotic genome (Keeling, 2004). Intestinal pathogen E. cuniculi is an obligate
intracellular parasite that infects broad spectrum of mammals including humans. It is an
important opportunistic pathogen in HIV infected patients. E. cuniculi genome possess only
1997 protein coding genes and according to molecular karyotype constructed by PFGE it is
organized as a diploid set of 11 chromosomes (Biderre et al., 1995; Katinka et al., 2001). In
the process of genome reduction, microsporidia underwent equally gene elimination, and
compaction that included the shortening of intergenic regions and genes themselves including
rDNA units. Encephalitozoon displays the highest gene density among eukaryotes and 80%
genes is shorter than their homologs in yeast (Biderre et al., 1995; Corradi et al., 2007;
12
Katinka et al., 2001; Keeling, 2004). With the exception of subtelomeric repeats at the ends of
chromosomes, genome of E. cuniculi is poor in repetitive elements that comprise large size
of genomes in most of eukaryotes and no transposones were detected in E. cuniculi genome.
Protein coding genes are localized in the GC rich core region of chromosomes that are flanked
by subtelomeric regions containing one rDNA unit (22 rDNA units in total) and repeats of
mini- and microsatellites (Biderre et al., 1995; Katinka et al., 2001).
1.2.3. Genome of Entamoeba histolytica (AMOEBOZOA)
Pathogenic amoeba E. histolytica causes amoebiasis that is significant cause of
morbidity and mortality in the Third World countries (Stanley, 2003). Genome of E.
histolytica (strain HM-1: IMSS) is 23 Mb large and 9938 genes coding proteins were
predicted. The genome is rich in introns that are present in 25% of genes (Loftus et al., 2005)
Chromosomes of E. histolytica do not condense and they are extremely fragile.
Together with the existence of various circular DNA molecules in the nucleus such as
plasmids bearing rRNA genes (200 copies per cell) (Willhoeft and Tannich, 1999) the
estimation of chromosome number is complicated and organism ploidy is still uncertain.
Different number of chromosomes was observed using different experimental methods.
Acridine orange staining revealed only six chromosomes (Gomez-Conde et al., 1998).
Separation by PFGE identified 31-35 chromosomes and suggested that E. histolytica is
tetraploid (Willhoeft and Tannich, 1999). The most recent study of E. histolytica
chromosomes by transmission electron microscopy suggests presence of linear 28-35 long
chromosomes that corresponds to PFGE analysis (Chavez-Munguia et al., 2006). Sizes of
corresponding chromosomes are variable in different isolated that is probably the result
of differences in the length of subtelomeric repeats, particularly of t-RNA arrays (Loftus et
al., 2005).
1.2.4. Genomes of Kinetoplastida (EXCAVATA)
Trypanosoma brucei
Subspecies of T. brucei are causative agents of African sleeping sickness in humans.
The majority of cases are caused by T. b. gambiense leading to chronic infection in Western
13
and Central Africa. T. b. rhodesiense is causative agent of acute sleeping sickness in Eastern
and Southern Africa (Malvy, 2011).
In the 26 Mb genome of T. brucei, 9068 genes were predicted. The genome is
characteristic by expansion of subtelomeric gene families (20% of the genome) involved in
antigenic variation of the parasite (Berriman et al., 2005). There are three classes of
chromosomes classified according to their size as minichromosomes (~100, 30 – 150 kb,
probably haploid), intermediate (1-5 chromosomes, 100 – 900 kb, probably haploid) and
megabase chromosomes (11 diploid chromosomes, 1 – 6 Mb). In kinetoplastids,
intermediate - and minichromosomes were found only in T. brucei where they comprise of
about 20% of total genome size. Megabase chromosomes contain housekeeping genes and
expression sites for metacyclic and bloodstream variants of surface glycoprotein (VSG)
genes. Variability in size observed among homologous chromosomes is in T. brucei caused
by length variation of VSG expression sites, telomeres and repetitive regions rich in
transposable elements. Intermediate chromosomes contain VSG expression sites and are
thought to originate by breakage of megabase chromosomes. Minichromosomes constitute
the reservoir of VSG genes. They also vary in size due to dynamics of their telomeres
(Ersfeld, 2011; Ersfeld et al, 1999). Another specific feature of T. brucei genome is the
presence of subtelomeric genes named retrotransposone hot spots that are clusters of intact
genes and pseudogenes with yet unknown function (Berriman et al. 2005; Coppel and Black,
2005).
Leishmania major
L. major causes cutaneous leishmaniasis. L. major Friedlin strain selected for genome
project is a user-friendly laboratory strain and model for study of immunological aspects of
leishmaniasis (Ravel et al., 1998). L. major genome size is ~34Mb, consists of 36
chromosomes analysed by PFGE and contains 8000 genes organized in polycistronic clusters
as it is typical for Kinetoplastida (Ivens et al., 2005; Ravel et al. 1998).
The genome is diploid or, more precisely, partially aneuploid for one or few
chromosomes (Sunkin et al., 2000). Typically for trypanosomatids, Leishmania
chromosomes do not condense in any stage of the cell division. Chromosomes of Leishmania
can not be identified by their size because of high size polymorphism between isolates and
species due to different length of subtelomeric repetitive sequences. Size polymorphism of
14
Leishmania homologous chromosomes is however not the result of recombination events like
in Plasmodium (Ravel et al, 1998; Sunkin et al., 2000).
1.2.5. Genome of Giardia intestinalis (Metamonada, EXCAVATA)
An intestinal parasite G. intestinalis causes diarrheal disease, giardiasis. Two major
genotypes assemblage A and B infect humans worldwide (Lalle, 2010). The genome of
Giardia is structurally compact, but to the less extend then in
Cryptosporidium or
Encephalitozoon (Morrison et al., 2007). Genome size determined for Giardia assemblages A
and B is about 11,7Mb and contain 4787 and 4691 protein - coding genes, respectively, with
minimum of introns and transposable elements (Franzén et al., 2009; Morrison et al., 2007).
Giardia has two nuclei that differ in DNA content and also chromosome number
(Tumova et al., 2007). Separation of giardial chromosomes by PFGE revealed five
chromosomes per haploid genome (Le Blancq and Adam, 1998). Chromosomes of G.
intestinalis are dynamic in the mean of their frequent rearrangment resulting in size
variation between homologs. This process involves particularly subtelomeric rRNA gene
units that are located on multiple chromosomes (Adam, 1992; Le Blancq and Adam, 1998;
Upcroft et al., 2005). Chromosomes of Giardia condense during mitosis and can be studied
by cytogenetic spreading technique. Giardia metaphase chromosomes are tiny (0,8 – 2,4 µm),
rod – shaped or ovoid structures without apparent constrictions and they can not be
distinguished based on their morphology. Chromosomes of Giardia consist of two sister
chromatids that are not linked together, as it is typical for eukaryotic chromosomes, probably
due to modification in cohesin protein complex (Tumova et al., 2007). Despite the absence of
constrictions, chromosomes of Giardia were characterized as monocentric with point
centromeres. It was deduced from the ultrastructural study of dividing cell showing the low
number of spindle microtubules attaching to the chromosomes in metaphase of mitosis and
characteristic immunostaining pattern of cenH3 histone variant, a marker protein of
centromeres (Dawson et al., 2007; Tumova et al., 2007).
Recent study by Tumova et al. (Tumova et al., 2007) revealed that Giardia is not
tetraploid with 10 chromosomes in each nucleus, as originally proposed (Le Blancq and
Adam, 1998), but the two nuclei possess different chromosome number and thus Giardia is
aneuploid for at least some chromosomes. Two dominant karyotypes were observed in
Giardia: i.) 9 + 11 chromosome pattern in isolates from genetic assemblage A and ii.) 10 + 11
chromosomes in isolates from assemblage B. This assymetry was found out to be stable.
15
1.2.6. Genomes of Trichomonas vaginalis and related species (Parabasala, EXCAVATA)
T. vaginalis is the causative agent of human trichomoniasis, the most widespread nonviral sexually transmitted disease worldwide (Conrad et al., 2011). Analysis of T. vaginalis
G3 genome revealed surprising large genome size: about 160 Mbp (Carlton et al., 2007). It is
the largest genome among sequenced parasitic prosists and with 60 000 protein coding
genes the genome has one of the highest coding capacities in eukaryotes. Several factors were
suggested to participace on the Trichomonas genome expansion.: (i) the genome underwent
several large scale genome duplications, (ii) specific gene families were amplified, (iii) the
genome has highly repetitive nature that was predicted to be 53,3% already in former study
of Wang and Wang (Wang and Wang, 1985). Various repetitive sequences such as
transposones or retrotransposones comprise about two-thirds (65%) of the genome (Carlton et
al., 2007). Several hundred copies of transposone mariner were identified here as the first
representative of the mariner family found in protists (Silva et al., 200ř).
T. vaginalis genome contains  250 rDNA units that are composed of genes coding
16S, 5,8S and 23S rRNAs. The units are arranged in classical head to tail tandem arrays. They
were mapped in a single nucleolar organizer region (NOR) by fluorescence in situ
hybridization (FISH) in the secondary constriction of chromosome IV (Carlton et al., 2007;
Zubacova et al., 2011). Previous studies of NORs in trichomonads based on silver staining
method failed to detect NOR in T. vaginalis, however, a single NOR was found in karyotypes
of related trichomonads Trichomitus batrachorum and Tritrichomonas augusta (Drmota and
Kral, 1997; Zubacova et al. 2011). Recently, FISH technique revealed single NOR also in
Tritrichomonas foetus and Trichomonas tenax, a sister taxon of T. vaginalis (TorresMachorro et al., 2009).
Karyotype of T. vaginalis as well as karyotypes of other trichomonads consists of
stable haploid set of chromosomes that condense during mitosis and can be studied using
spreading technique and other cytogenetic techniques such as C- and fluorescence-banding or
FISH. The culture of trichomonads can be enriched in cells arrested in metaphase by using
mitotic drug colchicine (Drmota and Kral, 1997; Yuh et al., 1998). In contrast to other
parasitic protists, numerous attempts for construction of molecular karyotype were
unsuccessful. Chromosomes of T. vaginalis can not be separated by PFGE probably due to
their large size.
16
Karyotype of T. vaginalis consists of six chromosomes that could be distiguish by
their size and morphology. They are tiny (1 – 2.4 µm) and consist of two sister chromatids
arranged in paralell, each with its own centromere. Genes encoding proteins of cohesin multi
– protein complex involved in sister chromatid cohesion, such as Rad21 are annotated in T.
vaginalis genome (Tumova et al., 2007; Zubacova and Tachezy, unpublished data). However,
sister chromatid cohesion as a thin connections between chromatids can be observed very
rarely on T. vaginalis metaphase microscopic preparations (Drmota and Kral,1997; Zubacova
unpublished observation).
Together with the examination of T. vaginalis, karyotypes,
species
including
genera
Tritrichomonas,
Tetratrichomonas,
related trichomonad
Pentatrichomonas,
Monocercomonas, Trichomitus and Hypotrichomonas were recently revised as available
analyses conducted between 1909 and 1931 were rather inconclusive (Drmota and Kral,
1997). Using spreading technique we described four to six chromosomes in their karyotypes.
The chromosome number was haploid and species-stable in all examined organisms
(Zubacova et al., 2008).
Cytogenetic techniques of chromosome examination including banding or FISH have
not been previously employed in study of trichomonad genome. Similarly to Giardia, our
attempts to visualize heterochromatin regions such as centromeres and AT- or GC- rich DNA
on T. vaginalis chromosomes using C – banding and fluorescence banding failed (Tumova et
al., 2007; Zubacova, unpublished). On the other hand, FISH technique was successfully
established for mapping of both multiple and single copy genes (Zubacova et al. 2011). This
technique was employed to map distribution of transposone mariner on T. vaginalis
chromosomes and in the study of genetic polymorhpism of different T. vaginalis isolates
using single – copy microsatellite markers (Carlton et al., 2007; Conrad et al., 2011).
The expansion of T. vaginalis genome might be associated with increased cell size. It was
hypothesized that large size of T. vaginalis genome and cell are related with pathogenicity of
this parasite, augmentation of its phagocytosis ability and reduction of its own ingestion by
host macrophages. Thus, genome sizes of eight selected trichomonad species of different
pathogenicity and phagocytic capacity have been studied. The estimation of nuclear DNA
content by flow cytometry revealed relatively large genomes (86 – 177 Mb) in all tested
species independently on their parasitic life style (Zubacova et al., 2008). Genome sequencing
of another trichomonad species is highly desired to elucidate parabasalid genome evolution
17
2.
Mitosis in parasitic protists
Mitosis is an essential process for cell life. Mitosis is a part of eukaryotic cell cycle
that includes chromosome segregation into daughter nuclei that proceed cell division. It is
divided into stages of profase, metaphase, anaphase and telophase that include changes in
chromosome movement and nuclear envelope dynamics. In a textbook mitosis, at prophase,
chromatin starts to condense to chromosomes that consist of two sister chromatids. The
chromatids are linked together via cohesin protein complex at the centromere. Microtubule
organizing center (MTOC) duplicates to form mitotic spindle and nuclear envelope
disintegrates. At metaphase, chromosomes are aligned at the center of the spindle to form
metaphase plate. They are anchored to spindle microtubules via multi-protein complexes
kinetochores that are built at centromeres one at each chromatid. Chromosome segregation
occurs at anaphase by the action of mitotic spindle and associated motor proteins. Sister
chromatids separate and begin to move towards opposite poles of the cell. At telophase,
nuclear envelope assemble around the daughter genomes, chromosomes decondense and cell
starts to divide into daughter cells (cytokinesis) (Gerald, 2011; Santaguida and Musacchio,
2009).
Every eukaryotic chromosome requires a centromere for proper segregation into
daughter cells. Centromeres evolved from telomeres during evolution of eukaryotic
chromosomes by breakage of ancestral circular chromosome and subsequent recovery of the
exposed DNA ends by retrotransposones followed by amplification of subtelomeric repeats
that gave rise to proto – centromeres. Phylogenetic analyses support the idea that all
centromeres, no matter how complex they are, share common ancestor based on repetitive
DNA derived from subtelomeric repeats (Villasante et al., 2007). In majority of eukaryotes
the centromere is not determined by specific DNA sequence motifs but epigenetically by
centromere specific proteins. Divergent variant of histone H3 generally called cenH3 (CENP
– A in mammals, Cid in Drosophila, Cse4 in yeast, CNA1 in Tetrahymena) is one and the
only marker of centromeres found in all studied eukaryotes with the only exception of
kinetoplastid protists (Lowell and Cross, 2004). CenH3s are essential for kinetochore
assembly on centromeres (Henikoff and Ahmad, 2005; Talbert and Henikoff, 2010).
Two types of chromosomes were described according to the structure of centromeres
and kinetochores. First, mononectric (or monokinetic) chromosomes have centromeres and
associated kinetochores localized in distinct chromosomal regions. These centromeres appear
as primary chromosomal constrictions. Monocentric chromosomes are present in majority of
18
model eukaryotes and are considered to be ancestral to holocentric chromosomes (Guerra et
al., 2010; Sagolla et al., 2006). In contrast, holocentric chromosomes do not have any
constrictions and their centromeres and so – called diffuse kinetochores are spread along
entire length of chromatids. This type of chromosomes was not studied so extensively in
comparison to „typical“ monocentric type despite they are relatively widespread. Holocentric
chromosomes occur commonly in invertebrates (for example in nematode worm
Caenorhabditis elegans), among protists in ciliates, and also in several green algae and plant
groups. Holocentric chromosomes were not observed in vertebrates (Dernburg, 2001; Guerra
et al., 2010; Mola and Papeschi, 2006; Tyler-Smith and Floridia, 2000).
Knowledge in the organization of crucial chromosome structures such as centromeres
and associated kinetochores is very limited in parasitic protists. Monocentric chromosomes
were described in Giardia intestinalis (Tumova et al., 2007) and Trichomonas vaginalis
(Drmota and Kral, 1997; Zubacova and Tachezy, unpublished data). Chromosomes of
Plasmodium falciparum, trypanosomatids and Entamoeba histolytica do not condense what
hampers to study their ultrastructure. From studies of centromeric DNA and kinetochores it is
unclear whether their chromosomes are monocentric or holocentric (Kelly et al., 2006; LopezRobles et al., 2000; Obado et al., 2007; Prensier and Slomianny, 1986). Centromeres were not
detected on Cryptosporidium chromosomes (Bankier et al., 2003).
Based on behavior of the nuclear envelope, two basic variants of mitosis can be
distinguished: open and closed. According to the symmetry of the spindle, mitosis is classified
as orthomitosis when the spindle has axial symmetry or pleuromitosis when excentric spindle
is present (Figure 1). Generally, higher eukaryotes undergo open mitosis. Mitotic spindle is
cytoplasmic and nuclear envelope disintegrates to allow association of spindle microtubules
with chromosome kinetochore (De Souza and Osmani, 2007). In eukaryotes with closed
mitosis, the nuclear envelope remains intact during whole cell cycle and mitotic spindle is
intranuclear. The organisms use the modification of nuclear pore complex or modify nuclear
transport to allow tubulin units enter the nucleus and assemble intranuclear spindle. Closed
mitosis occurs in unicellular eukaryotes such as fungi (e. g. yeasts) and trypanosomes (De
Souza and Osmani, 2007; Ersfeld and Gull, 1997; Raikov, 1994). In addition to open and
closed mitosis, variants of this process have evolved in various lineages of eukaryotes.
Extreme mitotic diversity occurs in protists however, details of this process are often poorly
understood (Brooks et al., 2011; Gerald, 2011; Raikov, 1994). However, along with gradual
release of genomic data, growing interest in investigation of molecular aspects of cell division
19
can be observed especially in parasitic protists with respect to developing antiparasitic drugs
that will be able to target cell division machinery and inhibit parasite´s reproduction.
Figure 1: Classification of mitosis based on nuclear envelope behavior and mitotic spindle localization (Raikov,
1994).
2.1. Mitosis in Apicomplexa
In Apicomplexa there are three types of cell division: i.) Endodyogeny is budding of
two daughter cells after mitosis within the mother cell. This type of reproduction is typical for
Toxoplasma tachyzoites (Dzierszinski et al., 2004). ii.) In schizogony repeated mitosis
without cytokinesis leads to multinucleated cells and subsequent generation of numerous
haploid progeny by cytokinesis. Schizogony occurs in Plasmodium life cycle and in slow
proliferating bradyzoites of Toxoplasma. iii.) Endopolygeny is characteristic for tissue
parasite Sarcocystis, where several rounds of genome duplication occur without nuclear
division leading to the large temporarily polyploid nucleus followed by budding into multiple
daughters (Figure 2). Multiple intranuclear spindles persist throughout the cell cycle
providing organization of chromosomes within polyploid nucleus (Brooks et al., 2011;
Dzierszinski et al., 2004; Striepen et al., 2007; Vaishnava et al., 2005).
20
Nuclear envelope persists during mitosis of Apicomplexa but there are pores within
the envelope for penetration of spindle microtubules. Two MTOCs (called also kinetic centers
and centriolar plaques in Plasmodium or centrocons in Toxoplasma and Sarcocystis ) are
extranuclear but are embedded within the outer side of the envelope giving rise to two bundles
of intranuclear spindle microtubules (Brooks et al., 2011; Gerald, 2011). This type of mitosis
is called semi open pleuromitosis.
Chromosomes do not condense in Apicomplexa. Throughout the cell cycle they
remain permanently attached to the mitotic spindles via kinetochores to maintain genome
integrity of polyploid stages. Chromosomes of Plasmodium do not align into central
metaphase plate. They have characteristic trilaminar kinetochores although majority of
conserved centromere – associated and kinetochoral components were not detected in the
genome that suggest the evolution of novelties in the protein composition of Plasmodium
kinetochore (Kelly et al., 2006; Prensier and Slomianny, 2006). Centromeres in Plasmodium
were defined on the base of DNA sequence motifs using etoposide – mediated topoisomerase
II cleavage assay. This assay is based on inhibition of activity of topoisomerase II by
anticancer grug etoposide. In metaphase topoisomerase II concentrates at the centromeres
playing role in sister chromatid decatenation. Etoposide blocks the activity of topoisomerase
II leaving doublestrand breaks in regions of centromeres and subsequent breakage of DNA at
the sites of centromeres (Brooks et al., 2011; Kelly et al., 2006). Plasmodium has „regional“
centromeres that are extremely AT rich (97%), they contain repetitive arrays and are
relatively small comprised of only 2 - 3 kb of DNA (the length of centromeric region
composed of repetitive satellite DNA in plants and metazoans can be up to Mb). Ortholog of
centromere marker protein cenH3 was identified in Plasmodium genome however its
chromosomal localization has not been studied yet (Kelly et al., 2006; Obado et al., 2007). In
contrast to Plasmodium, centromeres of Toxoplasma chromosomes do not share characteristic
DNA sequence motif and are the sites of accumulation of di – and trimethylated lysine 9 of
core histone H3 that is a typical modification of heterochromatin flanking the centromeres.
Centromeres of Toxoplasma were identified by immunolocalization of cenH3 marker histone
and etoposide – mediated topoisomerase II cleavage assay. Centromeres immunostaining also
showed that during mitosis, Toxoplasma chromosomes align to classical metaphase plate
located very close to envelope between two MTOCs, the centrocone. The centrocone together
with persisting spindle microtubules and probably proteins of nuclear envelope regulate
organization of chromosomes throughout the parasite development to ensure genome stability
during rapid reproduction (Brooks et al., 2011).
21
Figure 2: Mitosis in Apicomplexa inside the host cell: During endodyogony Toxoplasma completes whole cell
cycle after each round of DNA replication. Plasmodium (schizogony) and Sarcocystis (endopolygeny) omit
cytokinesis and/or nuclear division after several round of DNA replication resulting in multinucleated or
polyploid cells (nuclei in grey, centrosomes in red, parasite membrane in purple) (Striepen et al., 2007).
2.2. Mitosis in Kinetoplastida
Kinetoplastid parasites proliferate extensively in the digestive tract of their insect
vectors, in host´s bloodstream (Trypanosoma brucei), tissue cells (Trypanosoma cruzi) or
macrophages (Leishmania) (Berriman et al., 2005; Ivens et al., 2005; Tyler and Engman,
2001). Mitosis in kinetoplastids is closed with intact nuclear envelope and intranuclear mitotic
spindle (Ogbadoyi et al., 2000). Distinct MTOCs have not been observed despite presence of
genes encoding proteins such as γ – tubulin and centrins that function in MTOC (Berriman et
al., 2011; Ersfeld and Gull, 1997).
There are two types of chromosomes in trypanosomes, megabase chromosomes and
minichromosomes that do not visibly condense during the mitosis probably due to altered
structure and properties of their core histones (Ersfeld et al., 1999). Segregation of
Trypanosoma megabase chromosomes and minichromosomes differs markedly (Ersfeld and
Gull, 1997). Despite their number and size, it was found that minichromosomes are divided
into daughter cells with high fidelity (Alsford et al., 2001). Minichromosomes are segregated
by central pole - to - pole spindle microtubules. It is not known how they interact with the
spindle, however, lateral gliding model along spindle microtubules has been proposed for
their segregation (Ersfeld, 2011; Gull et al., 1998). Megabase chromosomes that have
trilaminar kinetochores are segregated by peripheral pole – to – kinetochore microtubules
22
(Figure 3). Interesting feature of Trypanosoma mitosis is that the number of large
chromosomes exceeds the number of kinetochores (11 pairs of megabase chromosomes
versus 10 trilaminar kinetochore – like structures) (Ersfeld and Gull, 1997; Ersfeld et al.,
1999; Gull et al., 1998). This could be explained by existence of kinetochore independent
mechanism of chromosomal segregation. Alternatively, some of the large chromosomes may
possess small and undetectable kinetochores like kinetochores of Sachcaromyces cerevisiae
that are defined functionally as well as biochemically but their ultrastructure is not visible by
electron microscopy. The numer of kinetochores do not corresponds to the chromosome
number not only in T. brucei but also in T. cruzi and Leishmania (Gull et al., 1998; Ogbadoyi
et al., 2000). The exact nature of kinetochores and centromeres of kinetoplastids is poorly
understood. Centromeric DNA has not been identified, although there are several sequence
candidates. Repetitive palindromes of 177 bp in T. brucei minichromosomes or 5,5 kb
repetitive region within chromosome I are suggested to have centromeric role (Lowell and
Cross, 2004). In Trypanosoma cruzi, centromeric DNA was mapped by chromosome
fragmentation. A 16 kb region required for chromosome mitotic stability is GC rich and
composed predominantly of degenerated retrotransposones, but lacks repetitive satellite DNA
typical for centromeres of higher eukaryotes. Clusters of subtelomeric repeats may have role
in maintenance of mitotic stability in Leishmania species (Obado et al., 2007).
Trypanosomes have poorly conserved and reduced components of centromeric and
core kinetochore protein complexes. For example, they lack the centromeric variant of histone
H3, cenH3 that was found in all eukaryotes studied to date ranging from those with small
point centromeres (yeast Saccharomyces cerevisiae) to those with holocentric chromosomes
(nematode C. elegans). Instead of cenH3, trypanosomes have telomere – specific H3V
histone variant that is conserved among kinetoplastids (Berriman et al., 2005; Lowell and
Cross, 2004). H3V immunostaining in combination with FISH revealed that H3V is localized
to telomeres of all classes of Trypanosoma chromosomes. Its function in telomeres is not
understood (Lowell and Cross, 2004). Protein sequence of Trypanosoma histone H3V shares
several common features with cenH3, such as divergent N – terminus and absence of
glutamine residue in α – 1 helix of the protein. Histone H3V, similarly as cenH3, associates
with the spindle during mitosis. Furthermore, centromeres are thought to evolved from
telomeres what is consistent with the observations that telomeres and subtelomeric regions in
several organisms have centromeric function (Lowell and Cross, 2004; Villasante et al. 2007).
For example, chromosome fragmentation experiments showed that satellite DNA of
subtelomeric regions of Leishmania major chromosomes have centromeric properties in
23
connection with maintaining chromosome stability (Dubessay et al., 2004). On the other hand,
unlike cenH3, histone H3V lacks N – terminal extension and insertion in loop 1 of the protein
structure. Moreover, H3V is not essential for chromosome segregation and cell viability,
while gene knock out of cenH3 lead to defects in mitosis so H3V is probably not functional
homolog of cenH3 (Regnier et al., 2005; Sanyal and Carbon, 2002).
Figure 3: Model of segregation of Trypanosoma large and minichromosomes during mitosis. Large
chromosomes (yellow) are segregated by kinetochore (green) dependent manner. Minichromosomes (red)
segregate by gliding along pole-to-pole spindle microtubules (blue) (Gull et al., 1998).
2.3. Mitosis in Microsporidia
Microsporidia divide by mitosis during their merogony and in the sporont stage inside
host´s cells. They divide by closed mitosis with intranuclear spindle that is a typical trait
of fungi. Chromosomes do not condense and chromatin is diffused inside the nucleus
(Bigliardi et al., 1998; De Souza and Osmani, 2007).
2.4. Mitosis in Entamoeba histolytica
E. histolytica trofozoites proliferate asexually in large intestine of vertebrate host by
unusual type of closed mitosis. Unlike in majority of eukaryotes, there is no evidence for cell
cycle checkpoint control in E. histolytica. Consequently, genome is repeatedly duplicated
(endoreduplication) without nuclear and cellular division that leads to polyploidy (Lohia,
2003). Mitosis of majority of eukaryotes with functional checkpoints is normally blocked
when it switches to endoreduplications. On the other hand, endoreduplication is also
developmental mechanism responsible for cell types differentiation and tissue growth. It is
widespread in plants where polyploidy can be detected in majority of plant tissues, while in
24
animals is limited to specific cell types (Kondorosi et al., 2000; Lee et al., 2009). Whether
endoreduplications have some biological significance in E. histolytica is not clear. Polyploid
amoebas occur in laboratory predominantly in media with depletion of nutrients (Lohia,
2003). It has been hypothesized that endoreduplication in E. histolytica represents an energy
saving mode of proliferation that does not require cytoskelet rearrangements and production
of new cell membranes or is used for effective nutrients uptake by increasing cell size (Lee et
al., 2009).
Molecular basis of Entamoeba chromosome segregation and nuclear division is still
not fully elucidated. E. histolytica chromosomes and their number are poorly defined despite
fluorescence and electron microscopy examinations. Chromosomes do not condense and
nuclear DNA is concentrated as a dense body called karyosome in the center of the nucleus.
Karyosome was considered to play a role in organizing of MTOC in early phases of mitosis.
Microtubules appear in Entamoeba cells only during mitosis, while they are not detectable
neither in nucleus nor in cytoplasm in interphase. Intranuclear mitotic spindle is unusually
arranged to rosettes of 7 – 12 microtubules. In dividing nucleus, 28 – 35 microtubular rosettes
were observed suggesting that the number of Entamoeba chromosomes is at least 35
considering that each rosette is associated with a single chromosome (Chavez-Munguia et al.,
2006).
No typical trilaminar kinetochores were observed on E. histolytica chromosomes. In
the only study of centromeres, CREST serum was tested in E.histolytica to detect
centromere/kinetochore regions in chromatin. CREST serum originates from human patients
suffering from autoimmune disease systemic scleroderma when antibodies against
centromeric and kinetochore proteins are produced (Fritzler et al., 2011; Lopez-Robles et al.,
2000). Among lower eukaryotes, CREST serum was successfully tested in Leishmania
mexicana chromosomes study (Lopez-Robles et al., 2000). In E. histolytica fixed trophozoites
CREST serum recognized 2 – 3 clusters of intranuclear round bodies considered by López –
Robles et al. (2000) to be centromeres. However, recognition sites of CREST antibodies in
Entamoeba chromatin were not studied in detail. Furthermore, the number of fluorescence
signals on microscopic preparations does not correspond with the number of chromosomes
(35). The nature of Entamoeba centromeres and kinetochores thus still remains a subject of
discussion and needs further study.
25
2.5. Mitosis in Giardia intestinalis
Binucleate trofozoites of G. intestinalis reproduce in the upper part of small intestine.
During the cell division, Giardia must duplicate not only nuclei but also multiple cytoskeletal
structures such as eight flagella, the ventral disc, the funis and the median body that occur in
and characterize trophozoites. Giardia employ semi – open mitosis (also described in related
diplomonad Hexamita inflata) with two extranuclear mitotic spindles. The giardia mitosis
includes all the phases that characterize mitosis of higher eukaryotes (Sagolla et al., 2006;
Tumova et al., 2007).
The mode of mitosis as well as presence of mitotic spindles were identified in Giardia
just recently by transmission electron microscopy (TEM) and 3D fluorescence deconvolution
microscopy. These studies revealed usual spindle microtubules based segregation of
chromosomes and disproved previous proposals of unconventional mode of genome
segregation based on spindle microtubule-independent mechanism (Solari et al., 2003).
During semi-open mitosis, nuclear membrane is retained, however, polar openings are
formed in the membrane and enable kinetochoral microtubules to enter the nucleus and
associate with chromosomes. Segregation of chromosomes proceeds in left – right axis of
bilaterally symmetrical trophozoite. Cytokinesis takes place along longitudinal axis of the
cell. Despite observed alignment of chromosomes into spindle midzone, no typical metaphase
plate was identified probably because of the small chromosome size or transient nature of this
step (Sagolla et al., 2006; Tumova et al., 2007).
Evidence of functionally conserved spindle in Giardia was based on behavior of
histone cenH3 in mitotic anaphase when it clusters at the leading edge of segregating DNA
towards spindle microtubule. Despite absence of direct evidence for attachment of
microtubules to kinetochores, the number of microtubules entering the nucleus seen on
electron microscopy preparations suggest that more than one microtubule is attached to one
kinetochore (Dawson et al., 2007; Sagolla et al., 2006). Conserved kinetochoral proteins were
not identified in G. intestinalis genome, however the presence of cenH3 histone variant, as an
essential basement for eukaryotic kinetochore assembly, indicates the existence of
kinetochoral structure (Dawson et al., 2007).
Giardia nuclei remain physically separated from each other during cell division
because of persistence of nuclear membrane that might lead to accumulation of allelic
heterozygosity. However, genome sequencing revealed remarkably low level of
heterozygosity between genome copies 0,01%. The mechanism of maintaining of
26
homozygosity of giardial nuclei is thus unknown. Nuclear fusions and meiotic recombinations
leading to reduction of heterozygosity were suggested, however, cytological evidence of such
events is missing (Morrison et al., 2007; Sagolla et al., 2006; Tumova et al., 2007).
2.6. Mitosis in Trichomonas vaginalis
The ultrastructure of T. vaginalis mitosis was studied by Brugerolle and colleagues
(Bricheux et al., 2007; Brugerolle, 1975). The phases of mitosis were described by Drmota
and Kral (Drmota and Kral, 1997) who found all typical mitotic phases. T. vaginalis
proliferates in urogenital tract of humans by a specific type of close mitosis named
cryptopleuromitosis (Brugerolle, 1975). Similarly to Giardia, trichomonads form
extranuclear mitotic spindle. However, polar opening were not observed and microtubules
interacts somehow with chromosomes at nuclear envelope (Raikov, 1994). As in other lower
eukaryotes with closed mitosis and extranuclear spindle, partial dissasembly and opening of
nuclear pore complexes of intact nuclear envelope can be considered to enable the direct
attachment of trichomonad spindle microtubules to kinetochores (De Souza and Osmani,
2007).
The mitotic spindle is assembled from two MTOCs called atractophores. Three types
of microtubules organize T. vaginalis extranuclear spindle: i.) pole to pole called
paradesmosis, ii.) pole to cytoplasm and iii.) pole to nucleus called kinetochoral microtubules
that attach to the outer side of nuclear membrane via a sort of dense material on the envelope
named by Brugerolle the kinetochore (Bricheux et al., 2007).
The structural basis of chromosomes segregation is poorly understood. Together with
mitotic spindle, the nuclear envelope is considered to play a role in T. vaginalis chromosome
segregation. It was suggested that the fluidity of nuclear membrane participates on movement
of chromosomes at anaphase (Raikov, 1994). Whether trichomonad chromosomes are
arranged along metaphase plate is not clear.
Proteins involved in T. vaginalis mitosis such as components of centromere and
kinetochore have not been studied experimentally. Genes coding for typical kinetochoral and
centromere - associated proteins are present in T. vaginalis genome (Zubacova and Tachezy,
unpublished data), however, their spectrum seems to be reduced in comparison to yeast and
higher eukaryotes. Inability to detect homologous components in T. vaginalis might also
reflect high diversity of Trichomonas sequences or presence of unique components of
27
kinetochore complex that remain to be described (Santaguida and Musacchio, 2009;
Zubacova and Tachezy, unpublished data).
Study of T. vaginalis centromeres is in its infancy. In initial study, we focused on
identification of centromeric variant of histone H3, cenH3. We identified three histone H3
variants and 20 core H3 histones. To identify putative cenH3, we examined nuclear
localization of all three H3 variants and a single core H3 as control using expression of
hemagglutinin-tagged proteins in T. vaginalis. One of the H3 variant appears to be a putative
cenH3 based on characteristic fluorescence pattern in nuclei. Immunostaining of TvcenH3 in
G1 phase nuclei revealed six dots that correspond to kinetochores on six chromosomes.
Twelve TvcenH3 fluorescence dots in doublets were observed in G2 phase of the cell cycle
after DNA replication. This pattern of cenH3 is characteristic for monocentric centromeres
appearance in interphase nuclei (Dernburg, 2001; Zubacova and Tachezy, unpublished data).
Our attempts to localize centromeres on Trichomonas vaginalis chromosomes using
CREST antibodies recognizing mammalian centromeric and kinetochore proteins were not
successful probably because of differences in protein sequences of major CREST antigens
such as CENP – A (mammalian cenH3 homolog) and CENP - B between humans and
trichomonads. Experiments with expression of heterologous cenH3 from G. intestinalis or
human CENP – A with the aim to identify the chromosomal position centromeres also failed
in Trichomonas. None of these proteins could functionally replace trichomonad cenH3 in
transfected trichomonads (Zubacova and Tachezy, unpublished data).
Unique localization of the second T. vaginalis H3 variant corresponded to histone
H3.3, a marker of transcriptionally active chromatin characterized by H3K4 methylation
(Dernburg, 2001; Talbert and Henikoff, 2010).
Despite completely different protein sequences, chromatin localization of the third T.
vaginalis H3 variant was identical with localization of core H3. Function of this H3 variant is
unknown (Zubacova and Tachezy, unpublished data).
28
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38
3. Aims

To compare nuclear DNA content and karyotypes in selected species of
Trichomonadea group.

To adapt fluorescence in situ hybridization (FISH) for mapping of repetitive sequences
and single copy genes on T. vaginalis chromosomes.

To map genes for ribosomal RNA, position and number of nucleolar organizer regions
on T. vaginalis chromosomes and genome distribution of transposone mariner.

To investigate histone H3 variants in T. vaginalis with focus on identification of
centromeric histone H3 (cenH3) as a marker protein of centromeres.
39
4. Publications
Carlton, J. M., Hirt R. P., Silva J. C., Delcher A. L., Schatz M., Zhao Q., Wortman J. R.,
Bidwell S. L., Alsmark U. C., Besteiro S., Sicheritz-Ponten T., Noel C. J., Dacks J.
B., Foster P. G., Simillion C., Van de Peer Y., Miranda-Saavedra D., Barton G.
J., Westrop G. D., Müller S., Dessi D., Fiori P. L., Ren Q., Paulsen I., Zhang H.,
Bastida-Corcuera F. D., Simoes-Barbosa A., Brown M. T., Hayes R. D.,
Mukherjee M., Okumura C. Y., Schneider R., Smith A. J., Vanacova S.,
Villalvazo M., Haas B. J., Pertea M., Feldblyum T. V., Utterback T. R., Shu C.
L., Osoegawa K., de Jong P. J., Hrdy I., Horvathova L., Zubacova Z., Dolezal P.,
Malik S. B., Logsdon J. M. Jr, Henze K., Gupta A., Wang C. C., Dunne R. L.,
Upcroft J. A., Upcroft P., White O., Salzberg S. L., Tang P., Chiu C. H., Lee Y.
S., Embley T. M., Coombs G. H., Mottram J. C., Tachezy J., Fraser-Liggett C. M.
and P. J. Johnson (2007). Draft genome sequence of the sexually transmitted
pathogen Trichomonas vaginalis. Science 315: 207-12.
Zubacova, Z., Z. Cimburek and J. Tachezy (2008). Comparative analysis of trichomonad
genome sizes and karyotypes. Molecular and Biochemical Parasitology 161: 49-54.
Conrad, M., Zubacova, Z., Dunn, L. A., Upcroft, J., Sullivan, S. A., Tachezy, J. and J.
M. Carlton (2011). Microsatellite polymorphism in the sexually transmitted human
pathogen Trichomonas vaginalis indicates a genetically diverse parasite. Molecular
and Biochemical Parasitology 175: 30-38.
Zubacova, Z., V. Krylov, and J. Tachezy (2011). Fluorescence in situ hybridization (FISH)
mapping of single copy genes on Trichomonas vaginalis chromosomes. Molecular
and Biochemical Parasitology 176: 135-37.
40
RESEARCH ARTICLE
Draft Genome Sequence of the
Sexually Transmitted Pathogen
Trichomonas vaginalis
Jane M. Carlton,1*† Robert P. Hirt,2 Joana C. Silva,1 Arthur L. Delcher,3 Michael Schatz,3
Qi Zhao,1 Jennifer R. Wortman,1 Shelby L. Bidwell,1 U. Cecilia M. Alsmark,2 Sébastien Besteiro,4
Thomas Sicheritz-Ponten,5 Christophe J. Noel,2 Joel B. Dacks,6 Peter G. Foster,7 Cedric Simillion,8
Yves Van de Peer,8 Diego Miranda-Saavedra,9 Geoffrey J. Barton,9 Gareth D. Westrop,4
Sylke Müller,4 Daniele Dessi,10 Pier Luigi Fiori,10 Qinghu Ren,1 Ian Paulsen,1 Hanbang Zhang,1
Felix D. Bastida-Corcuera,11 Augusto Simoes-Barbosa,11 Mark T. Brown,11 Richard D. Hayes,11
Mandira Mukherjee,11 Cheryl Y. Okumura,11 Rachel Schneider,11 Alias J. Smith,11
Stepanka Vanacova,11 Maria Villalvazo,11 Brian J. Haas,1 Mihaela Pertea,3 Tamara V. Feldblyum,1
Terry R. Utterback,12 Chung-Li Shu,13 Kazutoyo Osoegawa,13 Pieter J. de Jong,13 Ivan Hrdy,14
Lenka Horvathova,14 Zuzana Zubacova,14 Pavel Dolezal,14 Shehre-Banoo Malik,15
John M. Logsdon Jr.,15 Katrin Henze,16 Arti Gupta,17 Ching C. Wang,17 Rebecca L. Dunne,18
Jacqueline A. Upcroft,19 Peter Upcroft,19 Owen White,1 Steven L. Salzberg,3 Petrus Tang,20
Cheng-Hsun Chiu,21 Ying-Shiung Lee,22 T. Martin Embley,2 Graham H. Coombs,23
Jeremy C. Mottram,4 Jan Tachezy,14 Claire M. Fraser-Liggett,1 Patricia J. Johnson11
We describe the genome sequence of the protist Trichomonas vaginalis, a sexually transmitted human
pathogen. Repeats and transposable elements comprise about two-thirds of the ~160-megabase
genome, reflecting a recent massive expansion of genetic material. This expansion, in conjunction
with the shaping of metabolic pathways that likely transpired through lateral gene transfer from
bacteria, and amplification of specific gene families implicated in pathogenesis and phagocytosis of
host proteins may exemplify adaptations of the parasite during its transition to a urogenital
environment. The genome sequence predicts previously unknown functions for the hydrogenosome,
which support a common evolutionary origin of this unusual organelle with mitochondria.
richomonas vaginalis is a flagellated
protist that causes trichomoniasis, a common but overlooked sexually transmitted human infection, with ~170 million cases
occurring annually worldwide (1). The extracellular parasite resides in the urogenital tract of
both sexes and can cause vaginitis in women
and urethritis and prostatitis in men. Acute
infections are associated with pelvic inflammatory disease, increased risk of human immunodeficiency virus 1 (HIV-1) infection, and
adverse pregnancy outcomes. T. vaginalis is a
member of the parabasilid lineage of microaerophilic eukaryotes that lack mitochondria and peroxisomes but contain unusual organelles called
hydrogenosomes. Although previously considered to be one of the earliest branching eukaryotic
lineages, recent analyses leave the evolutionary
relationship of parabasalids to other major eukaryotic groups unresolved (2, 3). In this article,
we report the draft sequence of T. vaginalis, the
first parabasalid genome to be described.
Genome structure, RNA processing, and
lateral gene transfer. The T. vaginalis genome
sequence, generated using whole-genome shotgun methodology, contains 1.4 million shotgun
reads assembled into 17,290 scaffolds at ~7.2×
coverage (4). At least 65% of the T. vaginalis
genome is repetitive (table S1). Despite several
procedures developed to improve the assembly
T
(4), the superabundance of repeats resulted in a
highly fragmented sequence, preventing investigation of T. vaginalis genome architecture. The
repeat sequences also hampered measurement of
genome size, but we estimate it to be ~160 Mb
(4). A core set of ~60,000 protein-coding genes
was identified (Table 1), endowing T. vaginalis
with one of the highest coding capacities among
eukaryotes (table S2). Introns were identified in
65 genes, including the ~20 previously documented (5). Transfer RNAs (tRNAs) for all 20
amino acids were found, and ~250 ribosomal
DNA (rDNA) units were identified on small
contigs and localized to one of the six T. vaginalis
chromosomes (Fig. 1).
The Inr promoter element was found in ~75%
of 5′ untranslated region (UTR) sequences (4),
supporting its central role in gene expression (6).
Intriguingly, the eukaryotic transcription machinery of T. vaginalis appears more metazoan than
protistan (table S3 to S5). The presence of a
T. vaginalis Dicer-like gene, two Argonaute
genes, and 41 transcriptionally active DEADDEAH-box helicase genes suggests the existence of an RNA interference (RNAi) pathway
(fig. S1). Identification of these components raises
the possibility of using RNAi technology to manipulate T. vaginalis gene expression.
During genome annotation, we identified 152
cases of possible prokaryote-to-eukaryote lateral
www.sciencemag.org
SCIENCE
VOL 315
gene transfer (LGT) [tables S6 and S7 and
Supporting Online Material (SOM) text], augmenting previous reports of conflicting phylogenetic relationships among several enzymes (7).
The putative functions of these genes are diverse, affecting various metabolic pathways
(fig. S2) and strongly influencing the evolution
of the T. vaginalis metabolome. A majority (65%)
of the 152 LGT genes encode metabolic enzymes, more than a third of which are involved
in carbohydrate or amino acid metabolism
(Fig. 2). Several LGT genes may have been
acquired from Bacteroidetes-related bacteria,
which are abundant among vertebrate intestinal
flora (fig. S3).
Repeats, transposable elements, and genome expansion. The most common 59 repeat
families identified in the assembly (4) constitute
~39 Mb of the genome and can be classified as
(i) virus-like; (ii) transposon-like, including
~1000 copies of the first mariner element identified outside animals (8); (iii) retrotransposonlike; and (iv) unclassified (Table 2). Most of the
1
Institute for Genomic Research, 9712 Medical Research Drive,
Rockville, MD 20850, USA. 2Division of Biology, Devonshire
Building, Newcastle University, Newcastle upon Tyne NE1 7RU,
UK. 3Center for Bioinformatics and Computational Biology,
University of Maryland, College Park, MD 20742, USA.
4
Wellcome Centre for Molecular Parasitology and Division of
Infection and Immunity, Glasgow Biomedical Research Centre,
University of Glasgow, Glasgow G12 8TA, UK. 5Center for
Biological Sequence Analysis, BioCentrum-DTU, Technical
University of Denmark, DK-2800 Lyngby, Denmark. 6Department of Biological Sciences, University of Calgary, 2500
University Drive NW, Calgary, Alberta T2N 1N4, Canada.
7
Department of Zoology, Natural History Museum, Cromwell
Road, London SW7 5BD, UK. 8Department of Plant Systems
Biology, Flanders Interuniversity Institute for Biotechnology
(VIB), Ghent University, Technologiepark 927, B-9052
Ghent, Belgium. 9School of Life Sciences Research, University of Dundee, Dow Street, Dundee DD1 5EH, UK.
10
Department of Biomedical Sciences, Division of Experimental and Clinical Microbiology, University of Sassari,
Viale San Pietro 43/b, 07100 Sassari, Italy. 11Department of
Microbiology, Immunology, and Molecular Genetics, University of California, Los Angeles, CA 90095–1489, USA.
12
J. Craig Venter Joint Technology Center, 5 Research Place,
Rockville, MD 20850, USA. 13Children’s Hospital Oakland
Research Institute, 747 52nd Street, Oakland, CA 94609, USA.
14
Department of Parasitology, Faculty of Science, Charles
University, 128 44 Prague, Czech Republic. 15Department of
Biological Sciences, Roy J. Carver Center for Comparative
Genomics, University of Iowa, Iowa City, IA 52242–1324, USA.
16
Institute of Botany, Heinrich Heine University, 40225
Düsseldorf, Germany. 17Department of Pharmaceutical Chemistry, University of California, San Francisco, CA 94143–2280,
USA. 18Department of Microbiology and Parasitology, University
of Queensland, Brisbane, Queensland 4072, Australia.
19
Queensland Institute of Medical Research, Brisbane, Queensland 4029, Australia. 20Bioinformatics Center/Molecular Medicine Research Center, Chang Gung University, Taoyuan 333,
Taiwan. 21Molecular Infectious Diseases Research Center, Chang
Gung Memorial Hospital, Taoyuan 333, Taiwan. 22Medical
Genomic Center, Chang Gung Memorial Hospital, Taoyuan 333,
Taiwan. 23Strathclyde Institute of Pharmacy and Biomedical
Sciences, John Arbuthnott Building, University of Strathclyde,
Glasgow G4 0NR, UK.
*To whom correspondence should be addressed. E-mail:
[email protected]
†Present address: Department of Medical Parasitology, New
York University School of Medicine, New York, NY 10011,
USA.
12 JANUARY 2007
207
RESEARCH ARTICLE
59 repeats are present in hundreds of copies
(average copy number ~660) located on small
(1- to 5-kb) contigs, and each repeat family is
extraordinarily homogenous, with an average
polymorphism of ~2.5%.
The lack of a strong correlation between copy
number and average pairwise difference between
copies (fig. S4) suggests that a sudden expansion
of the repeat families had occurred. To estimate
the time of expansion, we compared the degree
of polymorphism among T. vaginalis repeats to
Table 1. Summary of the T. vaginalis genome
sequence data. Assembly size (bp, base pairs)
includes all contigs and differs from estimated
genome size of ~160 Mb (4). The scaffold size is the
minimum scaffold length, such that more than half
the genome is contained in scaffolds of at least that
length. The number of predicted genes may include
low-complexity repeats or novel transposable
elements rather than true T. vaginalis genes, but
in the absence of decisive evidence these remain in
the gene set. The number of evidence-supported
genes includes those with either similarity to a
known protein (E < 1 × 10–10, >25% length of
protein) or similarity to an expressed sequence tag
(>95% identity over >90% length of the gene). A
total of 763 rDNA fragments (258 copies of 28S,
254 copies of 18S, and 251 copies of 5.8S) were
identified.
Feature
Value
Genome
Size of assembly (bp)
176,441,227
G+C content (%)
32.7
No. of scaffolds
17,290
N50 scaffold size (bp)
68,338
Protein-coding genes
No. of predicted genes
59,681
No. of evidence-supported genes
25,949
No. of genes with introns
65
Mean gene length (bp)
928.6
Gene G+C content (%)
35.5
Gene density (bp)
2956
Mean length of intergenic
1165.4
regions (bp)
Intergenic G+C content (%)
28.8
Non–protein-coding genes
Predicted tRNA genes
479
Predicted 5.8S, 18S, and
~250
28S rDNA units
the divergence between T. vaginalis and its sister
taxon T. tenax, a trichomonad of the oral cavity
(9), for several protein-coding loci (4). Our results indicate that repeat family amplification
occurred after the two species split (table S8).
Several families have also undergone multiple
expansions, as implied by bi- or trimodal distribution of pairwise distances between copies
(fig. S5). T. vaginalis repeat families appear to be
absent in T. tenax but are present in geographically diverse T. vaginalis (4), consistent with the
expansion having occurred after speciation but
before diversification of T. vaginalis.
The large genome size, high repeat copy
number, low repeat polymorphism, and evidence
of repeat expansion after T. vaginalis and T. tenax
diverged suggest that T. vaginalis has undergone
a very recent and substantial increase in genome
size. To determine whether the genome underwent
any large-scale duplication event(s), we analyzed age distributions of gene families with
five or fewer members (4). A peak in the age
distribution histogram of pairs of gene families
was observed (fig. S6), indicating that the genome underwent a period of increased duplication, and possibly one or more large-scale
genome duplication events.
Metabolism, oxidative stress, and transport.
T. vaginalis uses carbohydrate as a main energy source via fermentative metabolism under aerobic and anaerobic conditions. We found the
parasite to use a variety of amino acids as energy
substrates (Fig. 2) (10), with arginine dihydrolase
metabolism a major pathway for energy production
(fig. S7) (11). We confirmed a central role for
aminotransferases (Fig. 2 and table S9) and
glutamate dehydrogenase as indicated previously
(12, 13); these pathways are likely catabolic but
may be reversible to allow the parasite to
synthesize glutamate, aspartate, alanine, glutamine, and glycine. Genes required for synthesis
of proline from arginine (fig. S7) and for threonine
metabolism (fig. S8) were identified. We also
identified a de novo biosynthesis pathway for
cysteine via cysteine synthase, an LGT candidate
(fig. S8) (14), and genes encoding enzymes involved in methionine metabolism, including its
possible regeneration (fig. S9).
Earlier studies indicated that de novo lipid
biosynthesis in T. vaginalis is confined to the
major phospholipid phosphatidylethanolamine
(PE) (15), whereas other lipids, including cholesterol, are likely acquired from exogenous
sources. We found an absence of several essential enzyme-encoding genes in the synthesis
and degradation pathways of nearly all lipids (4),
in contrast to the PE synthetic pathway, which
appears complete; however, experimental verification of these results is required.
T. vaginalis is microaerophilic with a primarily anaerobic life style and thus requires redox and
antioxidant systems to counter the detrimental
effects of oxygen. Genes encoding a range of defense molecules, such as superoxide dismutases,
thioredoxin reductases, peroxiredoxins, and rubrerythrins, were identified (table S10).
T. vaginalis demonstrates a broad range of
transport capabilities, facilitated by expansion of
particular transporter families, such as those for
sugar and amino acids (table S11). The parasite
also possesses more members of the cationchloride cotransporter (CCC) family than any
other sequenced eukaryote, likely reflecting
osmotic changes faced by the parasite in a mucosal
environment.
None of the proteins required for glycosylphosphatidylinositol (GPI)-anchor synthesis
were identified in the genome sequence, making T. vaginalis the first eukaryote known to
lack an apparent GPI-anchor biosynthetic pathway. Whether T. vaginalis has evolved an unusual biosynthetic pathway for synthesis of its
nonprotein lipid anchors, such as the inositolphosphoceramide of surface lipophosphoglycans
(16), remains to be determined.
Massively expanded gene families. Many gene
families in the T. vaginalis genome have undergone expansion on a scale unprecedented in unicellular eukaryotes (Table 3). Such “conservative”
gene family expansions are likely to improve an
organism’s adaptation to its environment (17).
Notably, the selective expansion of subsets of
the membrane trafficking machinery, critical
for secretion of pathogenic proteins, endocytosis of host proteins, and phagocytosis of bacteria and host cells (table S12), correlates well
with the parasite’s active endocytic and phagocytic life-style.
Massively amplified gene families also occur
in the parasite’s kinome, which comprises ~880
genes (SOM text) encoding distinct eukaryotic
protein kinases (ePKs) and ~40 atypical protein
Fig. 1. Karyotype and fluorescent in situ hybridization (FISH)
analysis of T. vaginalis chromosomes. (A) Metaphase chromosome squashes of T. vaginalis
reveal six chromosomes (I to VI).
(B) FISH analysis using an 18S
rDNA probe shows that all ~250
rDNA units localize to a single
chromosome. (C) In contrast, the
Tvmar1 transposable element
(8) is dispersed throughout the
genome.
208
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RESEARCH ARTICLE
kinases, making it one of the largest eukaryotic
kinomes known. The parasite has heterotrimeric guanine nucleotide–binding proteins and
components of the mitogen-activated protein kinase (MAPK) pathway, suggesting yeast-like
signal transduction mechanisms. Unusually,
the T. vaginalis kinome contains 124 cytosolic
tyrosine kinase–like (TKL) genes, yet completely
lacks receptor serine/threonine ePKs of the TKL
family. Inactive kinases were found to make up
17% of the T. vaginalis kinome (table S13); these
may act as substrates and scaffolds for assembly of
signaling complexes (18). ePK accessory domains
are important for regulating signaling pathways,
but just nine accessory domain types were
identified in 8% (72/883) of the T. vaginalis ePKs
(table S14), whereas ~50% of human ePKs contain at least 1 of 83 accessory domain types. This
suggests that regulation of protein kinase function
and cell signaling in T. vaginalis is less complex
than that in higher eukaryotes, a possible explanation for the abundance of T. vaginalis ePKs.
T. vaginalis possesses several unusual cytoskeletal structures: the axostyle, the pelta, and the
costa (19). Most actin- and tubulin-related components of the cytoskeleton are present (table S15),
with the exception of homologs of the actin motor
myosin. In contrast, homologs of the microtubular
motors kinesin and dynein are unusually abundant
(Table 3). Thus, T. vaginalis intracellular transport
mechanisms are mediated primarily by kinesin
and cytoplasmic dynein, as described for Dictyostelium and filamentous fungi, raising the possibility that the loss of myosin-driven cytoplasmic
transport is not uncommon in unicellular eukaryotes (20). Whether the structural remodeling of
amoeboid T. vaginalis during host colonization
(see below) is actin-based, as described for other
eukaryotes, or driven by novel cytoskeletal rearrangements remains an open question.
We identified homologs of proteins involved
in DNA damage response and repair, chromatin
restructuring, and meiosis, the latter a process not
thought to occur in the parasite (table S16). Of the
29 core meiotic genes found, several are general
repair proteins required for meiotic progression in
other organisms (21), and eight are meiosisspecific proteins. Thus, T. vaginalis contains either
recent evolutionary relics of meiotic machinery or
genes functional in meiotic recombination in an
as-yet undescribed sexual cycle.
Molecular mechanisms of pathogenesis.
T. vaginalis must adhere to host cells to establish
and maintain an infection. A dense glycocalyx
composed of lipophosphoglycan (LPG) (Fig. 3)
and surface proteins has been implicated in
adherence (22), but little is known about this
critical pathogenic process. We identified genes
encoding enzymes predicted to be required for
LPG synthesis (table S17). Of particular interest are the genes required for synthesis of an
unusual nucleotide sugar found in T. vaginalis
LPG, the monosaccharide rhamnose, which is
absent in the human host, making it a potential
drug target. Genes (some of which are LGT
candidates) were identified that are involved in
sialic acid biosynthesis, consistent with the reported presence of this sugar on the parasite
surface (23).
We identified eight families containing ~800
proteins (4) that represent candidate surface
molecules (Fig. 3 and table S18), including ~650
GLYCOLYSIS
arginine
16
NH 3
15
18
hydroxypyruvate
14
17
glycerate
24
3-phosphoglycerate
3-phospho- 25
hydroxypyruvate
phosphoserine
27
19
carbamoyl
phosphate
13
ornithine
5
11
NH3
26
CO 2
12
citrulline
urea
NH3
putrescine
2-phosphoglycerate
serine
20
22
glycine
CO 2
1-pyrroline5-carboxylate
21
threonine
proline
phosphohomoserine
NH 3
α-ketobutyrate
methanethiol,
NH3
31
methionine
pyruvate
3-mercaptopyruvate
alanine
9
3, 40
3
5
glutamate
2
NH3 , H2 S
tyrosine,
phenylalanine
tryptophan
indole,
2 NH
3
38
3
aspartate
34
S-adenosylmethionine
33
35
4
asparagine
lysine
6
31
homocysteine
32
methylthio adenosine
29
phosphoenol NH3 ,
pyruvate
H 2S
NH 3
H 2S
cysteine
9
30
22
36
cystine
NH3
23
10
PYRUVATE
METABOLISM
28
8
7
1
NH3
39
leucine,
isoleucine,
NH3 valine
NH 3
CO 2
NH 3
S-adenosylhomocysteine
glutamine
4-aminobutanoate
8-amino-7-oxononanoate
37
1-aminocyclopropane1-carboxylate
7,8-diaminononanoate
α-ketoglutarate
S-adenosyl-4-methylthio2-oxobutanoate
Fig. 2. Schematic of T. vaginalis amino acid metabolism. A complete description of enzymatic reactions
(represented as numbers) is given in the SOM text. Broken lines represent enzymes for which no gene
was identified in the genome sequence, although the activity would appear to be required. Green boxes
indicate enzymes encoded by candidate LGT genes.
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VOL 315
highly diverse BspA-like proteins characterized
by the Treponema pallidum leucine-rich repeat,
TpLLR. BspA-like proteins are expressed on the
surface of certain pathogenic bacteria and mediate
cell adherence and aggregation (24). The only
other eukaryote known to encode BspA-like
proteins, the mucosal pathogen Entamoeba histolytica (25), contains 91 such proteins, one of
which was recently localized to the parasite
surface (26).
There are >75 T. vaginalis GP63-like proteins,
homologs of the most abundant surface proteins of
Leishmania major, the leishmanolysins, which
contribute to virulence and pathogenicity through
diverse functions in both the insect vector and the
mammalian host (27). Most T. vaginalis GP63like genes possess the domains predicted to be
required for a catalytically active metallopeptidase,
including a short HEXXH motif (28) (Fig. 3).
Unlike trypanosomatid GP63 proteins, which are
predicted to be GPI-anchored, most T. vaginalis
GP63-like proteins have a predicted C-terminal
transmembrane domain as a putative cell surface anchor, consistent with the apparent absence of GPI-anchor biosynthetic enzymes.
Other T. vaginalis protein families share domains
with Chlamydia polymorphic membrane proteins, Giardia lamblia variant surface proteins,
and E. histolytica immunodominant variable
surface antigens (Fig. 3 and table S18).
After cytoadherence, the parasite becomes
amoeboid, increasing cell-to-cell surface contacts
and forming cytoplasmic projections that interdigitate with target cells (19). We have identified
genes encoding cytolytic effectors, which may be
released upon host-parasite contact. T. vaginalis
lyses host red blood cells, presumably as a means
of acquiring lipids and iron and possibly explaining the exacerbation of symptoms observed during
menstruation (29). This hemolysis is dependent on
contact, temperature, pH, and Ca2+, suggesting the
involvement of pore-forming proteins (30) that
insert into the lipid bilayer of target cells, mediating osmotic lysis. Consistent with this, we have
identified 12 genes (TvSaplip1 to TvSaplip12)
containing saposin-like (SAPLIP) pore-forming
domains (fig. S10). These domains show a predicted six-cysteine pattern and abundant hydrophobic residues in conserved positions while
displaying high sequence variability (fig. S11).
The TvSaplips are similar to amoebapore proteins secreted by E. histolytica and are candidate
trichopores that mediate a cytolytic effect.
The degradome. Peptidases perform many
critical biological processes and are potential virulence factors, vaccine candidates, and
drug targets (31). T. vaginalis contains an expanded degradome of more than 400 peptidases (SOM text), making it one of the most
complex degradomes described (table S19).
Of the three families of aspartic peptidases (table
S20), T. vaginalis contains a single member of
the HIV-1 retropepsin family that might serve as
a putative candidate for anti-HIV peptidase inhibitors. Many studies have implicated papain
12 JANUARY 2007
209
RESEARCH ARTICLE
Table 2. Summary of highly repetitive sequences in the genome of T. vaginalis. The 31 unclassified repeat
families have been collapsed into one group.
Repeat name
R128b
R169a
R1794
R299a
R3a
R9a
R947a
Average (total)
R8
R107
R119
R11b
R128a
R130a
R165
R178
R204
R210
R2375
R242b
R26b
R289a
R309b
R414a
R41b
R473
Integ.a95313
Average (total)
R1407.RT
copia.a38393
Average (total)
Average (total)
Total all
Average all
Putative identity
Copy
no.
Viral
203
2348
903
752
2243
1037
867
873
954
2637
831
663
518
669
931
1283
Transposon
Mariner transposase
982
1304
Integrase
384
2246
Mutator-like profile
282
2954
Integrase
1842
981
HNH endonuclease
200
800
Mutator-like profile
173
1129
Mutator-like profile
365
2410
Integrase
580
2433
Integrase
51
1323
Mutator-like profile
75
2127
Endonuclease
566
765
Integrase
49
1151
Integrase
1148
1141
Integrase
54
1250
Integrase
56
1601
Integrase
37
909
Integrase
927
1184
Integrase
19
1124
Integrase
68
1301
414
1481
Retrotransposon
Reverse transcriptase
6
2349
Copia-like profile
7
11001
6.5
6675
31 unclassified families
793
1131
Poxvirus D5 protein
Phage tail fiber prot
Hypothetical
KilA-N terminal domain
Poxvirus D5 protein
KilA-N terminal domain
KilA-N terminal domain
660
family cysteine peptidases as virulence factors
in trichomonads; we identified >40 of them, highlighting the diversity of this family. Cysteine
peptidases that contribute to the 20S proteosome
(ubiquitin C-terminal hydrolases) are abundant
(117 members, ~25% of the degradome), emphasizing the importance of cytosolic protein degradation in the parasite. T. vaginalis has nine
NlpC/P60-like members (table S20; several of
which are LGT candidates), which play a role in
bacterial cell wall degradation and the destruction
of healthy vaginal microflora, making the vaginal
mucosa more sensitive to other infections.
We also identified many subtilisin-like and
several rhomboid-like serine peptidases, candidates for processing T. vaginalis surface proteins. In addition to the first asparaginase-type
of threonine peptidase found in a protist, 13
families of metallopeptidases were also identi-
210
Length
(bp)
Cumulative
length (bp)
Average
pairwise
difference (%)
370,658
607,929
1,879,762
655,984
1,721,187
503,959
298,778
(6,038,257)
1.2
3.9
3.0
6.6
4.6
3.6
3.1
3.7
1,158,473
518,936
303,256
1,634,764
131,491
137,526
368,152
636,901
51,185
118,581
329,717
45,564
1,175,921
53,674
73,737
27,708
807,711
15,254
22,023
(7,610,574)
0.5
0.9
1.1
2.1
1.5
0.7
1.1
1.5
1.6
0.9
5.4
2.5
2.6
2.2
1.4
1.4
2.5
1.3
2.3
1.8
6,833
16,504
(23,337)
1.0
7.5
4.3
(24,617,704)
38,289,872
2.6
1450
2.5
fied, as well as three cystatin-like proteins, natural peptidase inhibitors, which may regulate
the activity of the abundant papain-like cysteine
peptidases (table S20).
The hydrogenosome. Several microaerophilic protists and fungi, including trichomonads
and ciliates, lack typical mitochondria and possess double-membrane hydrogenosomes, which
produce adenosine triphophate (ATP) and molecular hydrogen through fermentation of metabolic intermediates produced in the cytosol.
Although the origin of these organelles has been
controversial, most evidence now supports a common origin with mitochondria (3). Few genes
encoding homologs of mitochondrial transporters,
translocons, and soluble proteins were identified in the T. vaginalis genome (fig. S12),
suggesting that its hydrogenosome has undergone reductive evolution comparable to other
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Table 3. Large gene families in T. vaginalis. Only
gene families, or aggregates of gene families
mediating a given process, for which there are
more than 30 members and that have been
assigned a putative function are listed. (See SOM
text for subfamily organization.) The small guanosine triphosphatases (GTPases) are the sum of Rab
and ARF small GTPases only; all other GTPases are
shown in table S12. ABC, ATP-binding cassette;
MFS, major facilitator superfamily; MOP, multidrug/oligosaccharidyl-lipid/polysaccharide; AAAP,
amino acid/auxin permease.
Gene family (functional unit)
Members
Protein kinases
927
BspA-like gene family
658
Membrane trafficking: small GTPases
328
rDNA gene cluster
~250
Cysteine peptidase (clan CA, family C19) 117
Membrane trafficking: vesicle formation 113
ABC transporter superfamily
88
GP63-like (leishmanolysin)
77
MFS transporter family
57
MOP flippase transporter family
47
Cysteine peptidase (clan CA, family C1)
48
AAAP transporter family
40
Dynein heavy chain
35
P-ATPase transporter family
33
Serine peptidase (clan SB, family S8)
33
Membrane trafficking: vesicle fusion
31
protists whose mitochondrial proteomes are
reduced (e.g., Plasmodium). Because nuclearencoded hydrogenosomal matrix proteins are
targeted to the organelle by N-terminal presequences that are proteolytically cleaved upon import
(32) similar to mitochondrial precursor proteins, we screened the genome for consensus 5to 20-residue presequences containing ML(S/T/A)
X(1...15)R (N/F/E/XF), MSLX(1...15)R(N/F/XF), or
MLR(S/N)F (28) motifs. A total of 138 genes
containing putative presequences were identified,
67% of which are similar to known proteins, primarily ones involved in energy metabolism
and electron-transport pathways (fig. S13 and
table S21).
The production of molecular hydrogen, the
hallmark of the hydrogenosome, is catalyzed by
an unusually diverse group of iron-only [Fe]hydrogenases that possess, in addition to a
conserved H cluster, four different sets of functional domains (fig. S14), indicating that hydrogen
production may be more complex than originally
proposed. The pathway that generates electrons
for hydrogen (fig. S12) is composed of many
proteins encoded by multiple genes (tables S22
to S24). Our analyses extend the evidence that
T. vaginalis hydrogenosomes contain the complete machinery required for mitochondria-like
intraorganellar FeS cluster formation (33) and
also reveal the presence of two putative cytosolic auxiliary proteins, indicating that hydrogenosomes may be involved in biogenesis of
cytosolic FeS proteins. Some components of
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RESEARCH ARTICLE
Fig. 3. Structural organization of putative T.
vaginalis surface molecules involved in host
cell adherence and cytotoxicity (28). Candidate
surface protein families
(a to f) are depicted as
part of the glycocalyx
(gray shading), known
to be composed of LPG
(g). The number of proteins with an inferred
transmembrane domain
(red box) is indicated at
right, with the size of the
entire family shown in
parentheses. Substantial
length variation exists
between and within
families (proteins are
not drawn to scale). Additional information can
be found in table S18.
cell volume reported for protists (41). T. vaginalis
is also a highly predatory parasite that phagocytoses bacteria, vaginal epithelial cells, and host
erythrocytes (42) and is itself ingested by macrophages. Given these interactions, it is tempting
to speculate that an increase in cell size could have
been selected for in order to augment the parasite’s
phagocytosis of bacteria, to reduce its own phagocytosis by host cells, and to increase the surface
area for colonization of vaginal mucosa.
References and Notes
FeS cluster assembly machinery have also been
found in mitosomes (mitochondrial remnants)
of G. lamblia, supporting a common evolutionary origin of mitochondria, hydrogenosomes,
and mitosomes (3).
A new predicted function of hydrogenosomes revealed by the genome sequence is
amino acid metabolism. We identified two components of the glycine-cleavage complex (GCV),
L protein and H protein. Another component of
this pathway is serine hydroxymethyl transferase (SHMT), which in eukaryotes exists as
both cytosolic and mitochondrial isoforms. A
single gene coding for SHMT of the mitochondrial type with a putative N-terminal hydrogenosomal presequence was identified. Because
both GCV and SHMT require folate (fig. S12),
which T. vaginalis apparently lacks, the functionality of these proteins remains unclear.
The 5-nitroimidazole drugs metronidazole
(Mz) and tinidazole are the only approved drugs
for treatment of trichomoniasis. These prodrugs
are converted within the hydrogenosome to toxic
nitroradicals via reduction by ferredoxin (Fdx)
(fig. S12). Clinical resistance to Mz (Mz R) is estimated at 2.5 to 5% of reported cases and rising
(34) and is associated with decreases in or loss
of Fdx (35). We identified seven Fdx genes with
hydrogenosomal targeting signals (tables S21
and S25), the redundancy of which provides
explanations for the low frequency of MzR and
for why knockout of a single Fdx gene does not
lead to MzR (35). Our analyses also provide clues
to the potential mechanisms that clinically resistant parasites may use, such as the presence of
nitroreductase (NimA-like), reduced nicotinamide
adenine dinucleotide phosphate (NADPH)–
nitroreductase, and NADH-flavin oxidoreductase
genes (tables S23 and S26), which have been
implicated in MzR in bacteria (36, 37).
Summary and concluding remarks. Our investigation of the T. vaginalis genome sequence
provides a new perspective for studying the biology of an organism that continues to be ignored
as a public health issue despite the high number
of trichomoniasis cases worldwide. The discovery of previously unknown metabolic pathways,
the elucidation of pathogenic mechanisms, and
the identification of candidate surface proteins
likely involved in facilitating invasion of human
mucosal surfaces provide potential leads for the
development of new therapies and novel methods
for diagnosis.
The analysis presented here of one of the most
repetitive genomes known has undoubtedly been
hampered by the sheer number of highly similar
repeats and transposable elements. Why did this
genome expand so dramatically in size? We
hypothesize that the most recent common ancestor of T. vaginalis underwent a population
bottleneck during its transition from an enteric
environment (the habitat of most trichomonads)
to the urogenital tract. During this time, the decreased effectiveness of selection resulted in
repeat accumulation and differential gene family expansion. Genome size and cell volume are
positively correlated (38, 39); hence, the increased
genome size of T. vaginalis achieved through
rapid fixation of repeat copies could have ultimately resulted in a larger cell size. T. vaginalis
cell volume is greater than that of T. tenax and
related intestinal species Pentratrichomonas hominis (40) and T. gallinae (41), and it generally
conforms to the relationship of genome size to
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VOL 315
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EMBO J. 16, 3484 (1997).
33. R. Sutak et al., Proc. Natl. Acad. Sci. U.S.A. 101, 10368
(2004).
12 JANUARY 2007
211
34. G. Schmid et al., J. Reprod. Med. 46, 545 (2001).
35. K. M. Land et al., Mol. Microbiol. 51, 115 (2004).
36. D. H. Kwon et al., Antimicrob. Agents Chemother. 45,
2609 (2001).
37. H. K. Leiros et al., J. Biol. Chem. 279, 55840
(2004).
38. T. Cavalier-Smith, Ann. Bot. (London) 95, 147 (2005).
39. T. R. Gregory, Biol. Rev. Camb. Philos. Soc. 76, 65
(2001).
40. B. M. Honigberg, G. Brugerolle, in Trichomonads
Parasitic in Humans, B. M. Honigberg, Ed. (SpringerVerlag, New York, 1990), pp. 5–35.
41. B. J. Shuter, Am. Nat. 122, 26 (1983).
42. J. G. Rendon-Maldonado, M. Espinosa-Cantellano,
A. Gonzalez-Robles, A. Martinez-Palomo, Exp. Parasitol.
89, 241 (1998).
43. We are grateful to the Trichomonas research community
and M. Gottlieb for support and guidance during the duration
of this project. We thank M. Delgadillo-Correa, J. Shetty,
S. van Aken, and S. Smith for technical support; T. Creasy,
S. Angiuoli, H. Koo, J. Miller, J. Orvis, P. Amedeo, and E. Lee
for engineering support; W. Majoros and G. Pertea for data
manipulation; and S. Sullivan and E. F. Merino for editing.
Funding for this project was provided by the National
Institute of Allergy and Infectious Diseases (grants UO1
AI50913-01 and NIH-N01-AI-30071). The Burroughs
Wellcome Fund and the Ellison Medical Foundation provided
funds for Trichomonas community meetings during the
genome project. The Chang Gung T. vaginalis Systems
Biology Project was supported by grants (SMRPD33002 and
SMRPG33115) from Chang Gung Memorial Hospital and
Taiwan Biotech Company Limited. This whole genome
shotgun project has been deposited at DNA Data Bank of
Japan/European Molecular Biology Laboratory/GenBank
under the project accession AAHC00000000. The version
described here is the first version, AAHC01000000.
Supporting Online Material
www.sciencemag.org/cgi/content/full/315/5809/207/DC1
Materials and Methods
SOM Text
Figs. S1 to S14
Tables S1 to S26
References
Data
24 July 2006; accepted 14 November 2006
10.1126/science.1132894
REPORTS
Spectropolarimetric Diagnostics of
Thermonuclear Supernova Explosions
Lifan Wang,1,2* Dietrich Baade,3 Ferdinando Patat3
Even at extragalactic distances, the shape of supernova ejecta can be effectively diagnosed by
spectropolarimetry. We present results for 17 type Ia supernovae that allow a statistical study of the
correlation among the geometric structures and other observable parameters of type Ia supernovae.
These observations suggest that type Ia supernova ejecta typically consist of a smooth, central,
iron-rich core and an outer layer with chemical asymmetries. The degree of this peripheral
asphericity is correlated with the light-curve decline rate of type Ia supernovae. These results lend
strong support to delayed-detonation models of type Ia supernovae.
ifferent supernova (SN) explosion mechanisms may lead to differently structured
ejecta. Type Ia supernovae (SNe) have
been used as premier tools for precision cosmology. They occur when a carbon and oxygen
white dwarf reaches the Chandrasekhar stability
limit, probably because of mass accretion in a
binary system, and is disrupted in a thermonuclear explosion (1). Decade-long debates have
discussed how the explosive nuclear burning is
triggered and how it propagates through the
progenitor star (2–6). Successful models generally start with a phase of subsonic nuclear
burning, or deflagration, but theorists disagree
on whether the burning front becomes supersonic
after the earlier phase of deflagration. An
explosion that does turn into supersonic burning
is called delayed detonation (4). The resulting
chemical structures are dramatically different for
deflagration (5) and delayed-detonation models
(7). For a pure deflagration model, chemical
clumps are expected to be present at all velocity
layers that burning has reached (5). For delayed
detonation, the detonation front propagates
D
1
Physics Department, Texas A&M University, College
Station, TX 77843–4242, USA. 2Lawrence Berkeley National Laboratory, Berkeley, CA 94720, USA. 3European
Southern Observatory, Karl-Schwarzschild-Strasse 2, D-85748
Garching, Germany.
*To whom correspondence should be addressed. E-mail:
[email protected]
212
through the ashes left behind by deflagration,
burns partially burned or unburned elements
further into heavier elements, and erases the
chemically clumpy structures generated by deflagration.
The polarized emission from a supernova is
caused by electron scattering in its ejecta. It is
sensitive to the geometric structure of the ejecta
(8). Electron scattering in an asymmetric ejecta
would produce nonzero degrees of polarization
(8). In continuum light, normal SNe Ia are only
polarized up to about 0.3%, but polarization as
high as 2% is found across some spectral lines
(9–14). The polarization decreases with time and
vanishes around 2 weeks past optical maximum.
The low level of continuum polarization implies
that the SN Ia photospheres are, in general, almost
spherical. The decrease of polarization at later
times suggests that the outermost zones of the
ejecta are more aspherical than the inner zones.
Similarly, the large polarization across certain
spectral lines implies that the layers above the
photosphere are highly aspheric and most likely
chemically clumpy (12–15).
In this study, we report polarimetry of 17 SNe
Ia. These are all the SNe Ia for which we have
premaximum polarimetry. The observations
(Table 1) were collected with the 2.1-m Otto
Struve Telescope of the McDonald Observatory of
the University of Texas and with one of the 8.2-m
Table 1. Identification, observing date, phase (in days relative to optical maximum light), telescope
(McD indicates McDonald Observatory), intrinsic degree of polarization across the Si II 635.5-nm line,
Dm15, and references for Dm15 values. Numbers in parentheses indicate 1-s error multiplied by 100.
SN
Date of observation
Phase
Telescope
1996X
1997bp
1997bq
1997br
1999by
2001V
2001el
2002bo
2002el
2002fk
2003W
2004dt
2004ef
2004eo
2005cf
2005de
2005df
14 April 1996
7 April 1997
25 April 1997
20 April 1997
9 May 1999
25 February 2001
26 September 2001
18 March 2002
14 August 2002
6 October 2002
31 January 2003
13 August 2004
11 September 2004
20 September 2004
1 June 2005
6 August 2005
8 August 2005
–4.2
–5.0
–3.0
–2.0
–2.5
–7.3
–4.2
–5.0
–6.4
–5.5
–4.5
–7.3
–4.1
–5.9
–9.9
–4.4
–4.3
McD 2.1
McD 2.7
McD 2.1
McD 2.1
McD 2.1
VLT
VLT
VLT
VLT
VLT
VLT
VLT
VLT
VLT
VLT
VLT
VLT
12 JANUARY 2007
VOL 315
SCIENCE
m
m
m
m
m
www.sciencemag.org
PSiII
Dm15
References
0.50(20)
0.90(10)
0.40(20)
0.20(20)
0.40(10)
0.00(07)
0.45(02)
0.90(05)
0.72(09)
0.67(10)
0.64(10)
1.60(10)
1.10(30)
0.71(08)
0.44(05)
0.67(14)
0.73(05)
1.30(02)
1.29(08)
1.17(02)
1.09(02)
1.79(01)
0.73(03)
1.16(01)
1.34(03)
1.38(05)
1.19(05)
1.30(05)
1.21(05)
1.54(07)
1.46(08)
1.12(06)
1.41(06)
1.29(09)
(19, 21)
(19, 22)
(19, 22)
(19, 23)
(24)
(19, 25)
(19, 26)
(19, 27)
(19)
see text
see text
(28)
see text
(28)
(28)
see text
see text
Molecular & Biochemical Parasitology 161 (2008) 49–54
Contents lists available at ScienceDirect
Molecular & Biochemical Parasitology
Comparative analysis of trichomonad genome sizes and karyotypes
Zuzana Zubáčová a , Zdeněk Cimbůrek b , Jan Tachezy a,∗
a
b
Department of Parasitology, Faculty of Science, Charles University in Prague, Viničná 7, Prague 128 44, Czech Republic
Center of Flow Cytometry, Institute of Microbiology, Academy of Science of the Czech Republic, Prague 14200, Czech Republic
a r t i c l e
i n f o
Article history:
Received 22 December 2007
Received in revised form 2 June 2008
Accepted 7 June 2008
Available online 19 June 2008
Keywords:
Trichomonas
Genome size
Cell size
C-value
Karyotype
a b s t r a c t
In parasitic protists, the genome sizes range from 2.9 Mb in Encephalitozoon cuniculi to about 160 Mb in
Trichomonas vaginalis. The suprisingly large genome size of the former human parasite resulted from the
expansion of various repetitive elements, specific gene families, and possibly from large-scale genome
duplication. The reason for this phenomenon, as well as whether other trichomonad species have undergone a similar genome expansion, is not known. In this work we studied the genomes of nine selected
species of the Trichomonadea group. We found that each species has a characteristic karyotype with a
stable and haploid number of chromosomes. Relatively large genome sizes were found in all the tested
species, although over a rather broad range (86–177 Mb). The largest genomes were typically observed
in the Trichomonas and Tritrichomonas genera (133–177 Mb), while Tetratrichomonas gallinarum contains
the smallest genome (86 Mb). The genome size correlated with the cell volume, however, no relationship between genome size and the site of infection or trichomonad phagocytic ability was observed. The
data presented here provide primary information towards selecting a trichomonad species for future
large-scale sequencing to elucidate the evolution of unusual parabasalid genomes.
© 2008 Elsevier B.V. All rights reserved.
1. Introduction
Trichomonads are anaerobic flagellated protists that belong to
the Parabasala group. Most trichomonad species are the commensals of vertebrates and insects inhabiting the host intestinal tract
[1–3]. However, they also include important pathogens such as Trichomonas vaginalis and Tritrichomonas foetus that cause sexually
transmitted diseases in humans and cattle, respectively [4,5]. T.
vaginalis is the first parabasalid for which the complete genome
sequence was recently described [6]. This genome sequence analysis revealed that the size of the T. vaginalis genome is unusually
large, at about 160 Mb [6], which is approximately sixfold larger
than was originally estimated [7] and represents one of the largest
genomes among parasitic protists. The gene prediction revealed
about 60,000 putative genes, which are organized into 6 chromosomes [6,8,9]. The reason for such a genome expansion remains
unclear, however, it has been hypothesized that it is a rather recent
event associated with the transition of the T. vaginalis ancestor
from an enteric to urogenital tract environment [6]. This transition
included an increase in the cell volume, which may positively correlate with the DNA content [10]. It was further speculated that this
larger cell volume might increase its phagocytic potential as well as
∗ Corresponding author. Tel.: +420 22195 1811; fax: +420 224 919 704.
E-mail address: [email protected] (J. Tachezy).
0166-6851/$ – see front matter © 2008 Elsevier B.V. All rights reserved.
doi:10.1016/j.molbiopara.2008.06.004
surface area for the parasite’s interaction with the host cell tissue
[6], factors which are highly relevant to trichomonad pathogenicity.
To test these hypotheses and contribute to our knowledge of
trichomonad genome organization, we determined and compared
genome sizes, cell and nuclei sizes as well as karyotypes among
selected trichomonad species including T. vaginalis, Trichomonas
tenax, Tetratrichomonas gallinarum, Pentatrichomonas hominis,
Tritrichomonas foetus, Tritrichomonas augusta, Monocercomonas colubrorum, Trichomitus batrachorum and Hypotrichomonas acosta,
which represent parasites or commensals of the oral cavity, intestine and urogenital tract in various vertebrates, including humans
(Table 1).
2. Materials and methods
2.1. Organisms and cultivation
Axenic cultures of the following organisms were used in the
study: T. vaginalis strains G3 [American Type Culture Collection
(ATCC) PRA-98] and T1 (provided by J.-H. Tai, Institute of Biomedical Sciences, Taipei, Taiwan), T. tenax strain Hs-4:NIH (ATCC 30207),
T. foetus strains KV-1 (ATCC 30924) and LUB-1 MIP [11], T. augusta
strain T-37 (ATCC 30077), T. gallinarum strain M3 [12], P. hominis
strain GII (isolated from human by Gibora 1981, Slovak Republic),
M. colubrorum strain R183 [1], T. batrachorum strain BUB (isolated
from Bufo bufo by Kulda 1991, Czech Republic), H. acosta strain L3
50
Z. Zubáčová et al. / Molecular & Biochemical Parasitology 161 (2008) 49–54
Table 1
Characterization of selected trichomonad species
Species
Host
Habitat
Phagocytosis
Pathogenicity
Trichomonas vaginalis
Trichomonas tenax
Tritrichomonas foetus
Ttritrichomonas augusta
Tetratrichomonas gallinarum
Pentatrichomonas hominis
Monocercomonas colubrorum
Trichomitus batrachorum
Hypotrichomonas acosta
Human
Human
Cattle
Amphibian, reptile
Galliform and anseriform birds
Mammals including human
Reptile
Amphibian
Reptile, chelonian
Urogenital tract
Oral cavity, respiratory tract
Urogenital tract
Intestine
Intestine
Intestine
Intestine
Intestine
Intestine
Yes [6]
Yes [32]
Limited [19,33]
Not observed [34]
Yes [35]
Yes [36]
Not observed [37]c
Yes [37]
Yes [38]
Yes
Noa
Yes
No
No
Nob
No
No
No
a
b
c
Bronchopulmonary infections by T. tenax have been reported in patients with cancer or other lung diseases [16–18,39].
An unusual case of pulmonary infection caused by P. hominis has been reported [40].
Čepička, unpublished observation.
(ATCC 30069). All organisms were grown in Diamond’s TryptoneYeast extract-Maltose medium (TYM) at pH 6.2 for T. vaginalis and
pH 7.2 for the others, supplemented with 10% (v/v) heat-inactivated
horse serum (Gibco-BRL) and ferrous ammonium citrate [13]. Trichomonads isolated from humans, cattle and birds were cultivated
at 37 ◦ C, the other species at 25 ◦ C.
2.2. Chromosome preparation
Metaphase chromosomes from colchicine-treated cells were
obtained by a modified spreading technique as described in [8,14].
Briefly, trichomonads were grown to the mid-log phase until the
cell density reached 0.5 to 1.5 × 106 cells per ml, depending on
the trichomonad species. Colchicine was then added to 5 ml of culture to reach a final concentration of 0.5–1 mM and the cells were
incubated for 8 h. After incubation, the cells were harvested by centrifugation at 1000 × g for 10 min. The pellets were washed once
with phosphate-buffered saline (PBS) pH 7.3 and hypothonized in
75 mM potassium chloride (KCl) for 15–20 min at room temperature. The suspensions were centrifuged at 1000 × g 10 min and the
cells in pellets were fixed twice with freshly prepared Carnoy fixation (ethanol–chloroform–acetic acid in the rate of 6: 3: 1). First, the
cells were fixed for 2 h at 4 ◦ C in 10 ml of the fixative. Then the cells
were spin down at 1300 × g for 10 min and fixed in 0.5–1 ml of the
fixative overnight at 4 ◦ C. Aliquots of the final suspension in the fixative solution were dropped onto microscope slides from a height
of about 100 cm. The spreads were dried at 45 ◦ C for 1 h. Chromosomes were stained with 4 ,6-diamidino-2-phenylindole (DAPI)
(0.5 (g/ml) in Vectashield mounting medium (VectorLabs). Preparations were observed using an IX81 microscope, photographed with
an IX2-UCB camera and processed using CellR̂ software (Olympus).
The metaphase figures were inspected from at least five independent experiments. At least 50 metaphase figures per isolate were
evaluated.
2.3. Estimation of cell and nucleus volume and genome size
correlation
relationships between genome sizes and cell volumes, and nuclei
volumes.
2.4. Flow cytometry
Cells in the logarithmic growth phase were stained with Hoechst
33342 (Molecular Probes) (final concentration 5 (g/ml) for 30 min
at 37 ◦ C (T. vaginalis, T. tenax, T. foetus, T. gallinarum, P. hominis) or
at 25 ◦ C (M. colubrorum, T. batrachorum, T. augusta, H. acosta). The
relative fluorescence intensities from Hoechst-stained nuclei were
measured and analyzed in an LSR II cytometer (Becton Dickinson,
San Jose, USA). Before the measurement, propidium iodide (Sigma)
was added to the sample (final concentration 5 (g/ml) to stain the
non-living cells. Hoechst 33342 was excited at 406 nm, emission
was detected using a 450/50 filter; 561 nm was used for propidium
iodide excitation and emission was detected using a 610/20 filter.
The flow rate was about 500 fluorescent events per second in individual samples. Data analysis was performed using FlowJo 8.3.3.
software (TreeStar, Oregon, USA). At least 10,000 living cells were
monitored for each histogram. The cytometer was calibrated with
reference to the genome size of the T. vaginalis G3 strain [6].
3. Results
3.1. Karyotype analysis
The chromatin of all studied trichomonads that were arrested
during metaphase of mitosis by colchicine condensed into
well-structured chromosomes (Fig. 1). Constant number of chromosomes was observed in more than 95% figures in all species
using DAPI-stained metaphase preparations (Table 2). Observed
deviations (smaller number of chromosomes) in less than 5% of
figures were considered to be artifacts of spreading technique
[14,15]. All trichomonads displayed a single set of chromosomes,
Table 2
Chromosome numbers determined in nine trichomonad species
Species
The dimensions of cells and nuclei were measured from digital
microphotographs of living cells using an Olympus BX51 microscope equipped with an IX2-UCB camera. The cells were attached
to microscope slides that were covered with poly-l-lysine (Sigma),
and observed using a combination of differential interference
contrast illumination and fluorescence of Hoechst 33342-stained
nuclei. Fifty non-dividing cells of each strain were measured using
the program QuickPHOTO MICRO 2.2 (Promicra). Approximate cell
and nucleus volumes (V) were calculated according to the formula
V = 3/4 ab2 , where “a” is the cell length, and “b” is the cell width.
Least-squares linear regression analyses were performed to test for
Trichomonas vaginalis
Trichomonas tenax
Tetratrichomonas gallinarum
Pentatrichomonas hominis
Tritrichomonas foetus
Tritrichomonas augusta
Monocercomonas colubrorum
Trichomitus batrachorum
Hypotrichomonas acosta
Chromosome number
This studya
Published data
6
6
5
6
5
5
4
6
5
6 [8,9]
3–6 [41]
Not available
5–6 [42]
5 [14]
5 [43], 4–8 [44]
Not available
6 [42,45], 4–5 [46], 4–8 [44]
Not available
a
The metaphase chromosome spreads from colchicine-treated cells were stained
with DAPI and counted using fluorescence microscopy.
Z. Zubáčová et al. / Molecular & Biochemical Parasitology 161 (2008) 49–54
51
Fig. 1. Karyotypes of representative species from Trichomonadidae [(A) Trichomonas vaginalis, (B) Trichomonas tenax, (C) Pentatrichomonas hominis, (D) Tetratrichomonas
gallinarum], Tritrichomonadidae [(E) Tritrichomonas augusta, (F) Tritrichomonas foetus], Monocercomonadidae [(G) Monocercomonas colubrorum, (H) Hypotrichomonas acosta],
and Trichomitidae [(I) Trichomitus batrachorum] groups. In metaphase of mitosis, only haploid chromosome sets were observed in all species. Each chromosome consists of
two sister chromatides. Arrows indicate chromatide constrictions. Scale bar is 10 ␮m.
each with two chromatids (Fig. 1). The chromatids were organized
in parallel with one or two constrictions observed in some pairs
of chromatids in each of the species examined. Although the
chromosomes were generally minute (0.9–2.4 ␮m in T. vaginalis),
they differed in morphology and size, producing a specific characteristic pattern for each species (Fig. 1 and Table 2). Homologous
chromosome pairs, the tetrads typically found in the first meiotic
division of other eukaryotes, were not observed in our specimens.
These results indicated that all selected species possess a stable,
haploid chromosome set.
3.2. Genome sizes
Trichomonad cells in the G1-phase of the cell cycle were used
to estimate DNA content by means of flow cytometry (Fig. 2). The
nuclei were stained with the DNA-specific fluorescent dye Hoechst
33342. At this stage, the DNA was organized as fine granules,
which were distributed homogenously within the nuclei except
the sites of nucleoli (Fig. 2). The plasma membrane of all trichomonad species used in the study was highly permeable to this
dye, no autofluorescence was observed (data not shown). T. vaginalis, the genome of which is completely sequenced [6], was used
as a standard for calibrating the cytometer. The other protists with
known genome sizes (Saccharomyces cerevisiae, Eimeria tenella, Giardia intestinalis and Trypanosoma brucei) appeared to be unsuitable
for the calibration due to smaller sizes of their genomes and/or
differences in nucleotide preferences (data not shown).
Fluorescence intensities corresponding to DNA content revealed
relatively large genomes in all nine trichomonad species compared to the genomes of other parasitic protists (Table 3). The
largest genome sizes were determined in the species of the Trichomonas and Tritrichomonas genera, ranging from 133 to 177 Mb.
The genomes of species from the other trichomonad genera displayed considerably smaller genomes, ranging from 86 to 114 Mb.
52
Z. Zubáčová et al. / Molecular & Biochemical Parasitology 161 (2008) 49–54
Fig. 2. Measurement of nuclear DNA amount (genome size) in trichomonads by flow
cytometry. (A) Histogram of fluorescence intensity corresponding to Hoechst 33342stained DNA in living cells of Tritrichomonas foetus. The G1-phase population was
used for genome size estimation and (B) Hoechst 33342-stained nuclei of G1-phase
Tritrichomonas foetus.
The smallest genome was found in the bird commensal T. gallinarum, which is almost 100 Mb smaller than the largest genome,
belonging to the cattle parasite T. foetus.
3.3. Correlation between genome sizes and sizes of cells and
nuclei
Published data concerning trichomonad cell sizes were determined by various methods, which made direct comparison
between trichomonad species difficult. Thus, we first determined
the sizes of trichomonad cells and their nuclei under standard conditions and calculated their approximate volumes (Table 4). The
largest cells as well as the largest nuclei were exhibited by the
Table 3
Genome sizes (C-values) determined in studied trichomonad species by means of
flow cytometry
Species, strain
Genome size (Mb) mean ± S.D. (n)b
Trichomonas vaginalis, G3
Trichomonas vaginalis, T1
Trichomonas tenax, Hs-4
Tritrichomonas foetus, KV-1
Tritrichomonas foetus, LUB
Tritrichomonas augusta, T-37
Tetratrichomonas gallinarum, M3
Pentatrichomonas hominis, GII
Monocercomonas colubrorum, R183
Trichomitus batrachorum, BUB
Hypotrichomonas acosta, L3
158–166a
154 ± 9 (8)
133 ± 4 (3)
177 ± 11 (4)
177 ± 12 (4)
165 ± 6 (3)
86 ± 10 (4)
94 ± 8 (3)
114 ± 17 (4)
125 ± 11 (3)
114 ± 16 (4)
Species
Genome size (Mb)
Toxoplasma gondii
Trypanosoma brucei
Plasmodium falciparum
Giardia intestinalis
Encephalitozoon cuniculi
80 [47]
35 [48]
23 [49]
12 [15]
2.9 [50]
Known genome sizes of selected parasitic protists are included for comparison.
a
Trichomonas vaginalis G3 strain with genome size value of 160 Mb (the precise
genome size was reported as a range from 158–166 Mb) [6] was used as an external
standard for calibration of the flow cytometer.
b
S.D.: standard deviation; n: number of independent experiments.
Fig. 3. Correlation between trichomonad genome sizes (Mbp) and cell and nucleus
volumes (␮m3 ). Genome sizes were positively correlated with cell volumes (R2 = 0.5)
as well as nucleus volumes (R2 = 0.8) in all tested species. The numbers in the plot
correspond to genome sizes.
species of the Trichomonas and Tritrichomonas genera, which correspond to the presence of the largest genomes (Table 3). Similarly, P.
hominis with the smallest cell and nucleus size possesses one of the
smallest genomes. Genome sizes were positively correlated with
cell volumes (R2 = 0.5) as well as nuclei volumes (R2 = 0.8) (Fig. 3).
4. Discussion
The data presented in this study revealed considerably large
genomes in all the selected trichomonad species in comparison to
other parasitic protists (Table 3). The genomes were organized into
four to six chromosomes forming a stable characteristic karyotype
for each species. The difference in published chromosome numbers for some species (Table 2) likely attributed to the much lower
limit of resolution of light microscopy available for distinguishing
the small trichomonad chromosomes in these studies, conducted
between 1909 and 1931.
The sizes of the genomes ranging from 86 to 177 Mb were neither directly related to the trichomonad’s pathogenicity, nor their
phagocytic potential, nor the site of the infection. Although the
largest genomes were found in the pathogenic species T. vaginalis and T. foetus, which inhabit the urogenital tract of the host
(160–177 Mb), a comparable genome size was obtained for T.
augusta (165 Mb), which is a commensal of the reptile intestine.
In addition, a relatively large genome was also found in T. tenax
(133 Mb), which is a commensal of the human oral cavity, although
its role as an etiologic agent in extraoral infections cannot be
excluded [16–18]. As far as phagocytic potential is concerned, tritrichomonads are not able to ingest host cells and only limited phagocytosis of cell debris was recently observed [19], although their cell
volume as well as genome size is comparable with Trichomonas
species, for which phagocytosis is a characteristic mechanism for
importing external material including whole bacteria, erythrocytes
or leukocytes [20]. In other words, the largest genomes were found
Z. Zubáčová et al. / Molecular & Biochemical Parasitology 161 (2008) 49–54
53
Table 4
Cell dimensions and approximate volumes of selected trichomonad species
Species
Cell
Nucleus
Length (␮m)
mean ± S.D.
Trichomonas vaginalis
Trichomonas tenax
Tritrichomonas foetus
Tritrichomonas augusta
Tetratrichomonas gallinarum
Pentatrichomonas hominis
Hypotrichomonas acosta
Monocercomonas colubrorum
Trichomitus batrachorum
14.8
14.2
13.28
16.71
12.01
6.92
11.2
8.82
12.38
±
±
±
±
±
±
±
±
±
1.6
1.92
2
2.39
0.92
0.92
1.54
2.08
1.58
Width (␮m)
mean ± S.D.
9.8
10
9.15
11.8
8.58
4.53
7.99
5.92
8.69
±
±
±
±
±
±
±
±
±
1.0
1.79
1.43
1.94
0.93
0.72
1.09
1.17
0.98
a
3
Volume (␮m )
Length (␮m)
mean ± S.D.
3.3
3.3
2.7
5.4
2
0.3
1.7
0.7
2.2
5.63
4.32
5.19
5.25
3.13
2.41
3.05
2.88
3.58
±
±
±
±
±
±
±
±
±
0.59
0.51
0.68
0.87
0.46
0.42
0.40
0.55
0.42
Width (␮m)
mean ± S.D.
3.00
2.56
3.47
3.58
2.47
1.82
1.97
1.89
2.78
±
±
±
±
±
±
±
±
±
0.50
0.44
0.41
0.60
0.32
0.38
0.30
0.42
1.50
Volumea (␮m3 )
0.12
0.06
0.15
0.16
0.04
0.02
0.03
0.02
0.07
Average of 50 specimens ± standard deviation.
a
The cell volume was calculated according to the formula V = 3/4 ab2 ; “a” is the cell length, “b” is the cell width.
in all the tested species of Trichomonas and Tritrichomonas genera,
without an apparent relationship to their parasitic style of life.
Next we considered the sizes of the genomes and phylogenetic
relationships between the studied trichomonads. As is apparent
from Fig. 4, both Trichomonadinae and Tritrichomonadinae groups
include genera from both ends of the genome size range. For example, P. hominis and T. gallinarum displayed considerably smaller
genomes, although they belong to the Trichomonadinae group
together with Trichomonas species. Similarly, M. colubrorum displayed a considerably smaller genome than Tritrichomonas species,
although M. colubrorum was shown to be closely related to T. foetus in various phylogenetic reconstructions [1,21]. Therefore, this
genome amplification (or reduction) may have happen independently within Trichomonadinae and Tritrichomonadinae groups.
The only significant correlation observed was between trichomonad genome sizes and the overall volumes of the cells as
well as their nuclei. Such a correlation was reported in all groups of
organisms from protists to vertebrates, although the exact causes
of this phenomenon are not clear [10]. It has been shown, however,
that an increase in a haploid genome (C-value) of large cells correlates with an increase in the amount of non-coding DNA, including
introns, pseudogenes and transposable elements, while a number
of genes remain comparable over a broad taxonomic scale. Consequently, an increase in non-coding DNA leads to a discrepancy
between the genome size and gene number, a phenomenon which
is also known as the C-value paradox [22,23]. For example in the
human genome, the protein coding genes comprise only 1.5% of
the genome, while the contribution of introns is over 25% [24].
Interestingly, introns are not involved in the expansion of the T.
vaginalis genome, in which only 65 introns were identified [6]. In
this organism, the genome expansion mainly involved the amplification of repeats and transposable elements, massive expansion of
certain gene families, and possibly large-scale genome duplications
[6]. The last two events may led to a multiplication of the majority of genes, which is reflected by the unusual high gene number,
reaching 60,000 in the T. vaginalis genome.
The unusual level of the gene multiplicity in the trichomonad
genome and large genome sizes might be related to the unusual
evolutionary history of the Parabasala group. Previously, monocercomonads were suggested to be an ancient parabasalid lineage
based on their morphological simplicity, from which trichomonads with a more complex karyomastigont (the organellar system,
which includes a nucleus attached to a mastigont consisting of flagella, central cytoskeletal structures, and a parabasal body) have
evolved [3,25]. This simple-to-complex scenario, however, was not
supported by molecular phylogeny and the absence of various
cytoskeletal structures in monocercomonads was most likely the
result of multiple loss in several trichomonad clades [1]. Suprisingly, several recent phylogenetic studies placed trichonymphids,
a group of organisms with very complex cytoskeletal structures,
at the root position of parabasalids [26–28] (Fig. 4). These organisms, living in the digestive tract of termites and wood-feeding
cockroaches, are characterized by the large cell size (about 100
times larger than species of the Trichomonadida group [3,29]), the
Fig. 4. Draft of phylogenetic relationships within Parabasala group [21,26–28]. The taxa included in this study are shown in bold.
54
Z. Zubáčová et al. / Molecular & Biochemical Parasitology 161 (2008) 49–54
multiple mastigonts, and the large nucleus with large number of
chromosomes [3,29]. For example, the nucleus of Trichonympha
major is about 17.6 ␮m in transversal diameter with 24 haploid
chromosomes [30]. As the karymastigont reproduces as a unit
structure [31], it is tempting to speculate that the trichonymphid
ancestor originally possessed complete multiple karyomastigonts
including nuclei as observed for example in Calonymphidae [3,29],
and later the multiple nuclei were transformed or somehow contributed to the formation of a single large hypermastigid nucleus
with presumably multiplicated genome. If the ancestral position
of trichonymphids is correct, we can further speculate that the
genome sizes observed in the studied trichomonad species might
result from a regressive evolution, which includes DNA content
decrease, simplification of the mastigont and decrease in cell size.
To elucidate the evolution of the parabasalid genome, more work
on large-scale genomics is required. In particular, it would be interesting to compare the genomes or selected genome markers in
organisms with total DNA contents at opposite ends of the spectrum. Based on our results, P. hominis seems to be an interesting
trichomonad species for such studies: (i) this organism displayed
one of the smallest genome in the range of selected trichomonad
species (94 Mb), in which less repeates and transposable elements
might be expected, (ii) it is living in the host intestine, an original environment from which an ancestor of T. vaginalis most likely
evolved, thus comparison with T. vaginalis genome may provide
new insight concerning events that were responsible for genome
size changes associated with transition of trichomonads from an
enteric environment to the urogenital tract [6] and (iii) as P. hominis is mostly harmless organism, the comparison with T. vaginalis
genome may help to pinpoint the gene families whose amplification
is related to T. vaginalis pathogenicity.
Acknowledgements
This work was supported by Czech Ministry of Education, Youth
and Sport, project MSM0021620858 and LC07032 (J.T.), and the
Grant Agency of Charles University 180/2006/B-BIO/PrF (Z.Z.).
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Molecular & Biochemical Parasitology 175 (2011) 30–38
Contents lists available at ScienceDirect
Molecular & Biochemical Parasitology
Microsatellite polymorphism in the sexually transmitted human pathogen
Trichomonas vaginalis indicates a genetically diverse parasite夽
Melissa Conrad a , Zuzana Zubacova b , Linda A. Dunn c , Jacqui Upcroft c , Steven A. Sullivan a ,
Jan Tachezy b , Jane M. Carlton a,∗
a
b
c
Department of Medical Parasitology, New York University Langone Medical Center, New York, NY 10010, USA
Department of Parasitology, Faculty of Science, Charles University in Prague, Vinicná 7, Prague 128 44, Czech Republic
Queensland Institute of Medical Research, Brisbane, QLD 4006, Australia
a r t i c l e
i n f o
Article history:
Received 3 July 2010
Received in revised form 19 August 2010
Accepted 23 August 2010
Available online 9 September 2010
Keywords:
Population genetics
Trichomonas vaginalis
Sexually transmitted infection
Microsatellite
FISH
Genetic markers
a b s t r a c t
Given the growing appreciation of serious health sequelae from widespread Trichomonas vaginalis infection, new tools are needed to study the parasite’s genetic diversity. To this end we have identified and
characterized a panel of 21 microsatellites and six single-copy genes from the T. vaginalis genome, using
seven laboratory strains of diverse origin. We have (1) adapted our microsatellite typing method to
incorporate affordable fluorescent labeling, (2) determined that the microsatellite loci remain stable in
parasites continuously cultured for up to 17 months, and (3) evaluated microsatellite marker coverage of
the six chromosomes that comprise the T. vaginalis genome, using fluorescent in situ hybridization (FISH).
We have used the markers to show that T. vaginalis is a genetically diverse parasite in a population of
commonly used laboratory strains. In addition, we have used phylogenetic methods to infer evolutionary
relationships from our markers in order to validate their utility in future population analyses. Our panel
is the first series of robust polymorphic genetic markers for T. vaginalis that can be used to classify and
monitor lab strains, as well as provide a means to measure the genetic diversity and population structure
of extant and future T. vaginalis isolates.
© 2010 Elsevier B.V. All rights reserved.
1. Introduction
Trichomoniasis, caused by the parasitic protist Trichomonas
vaginalis, is the most prevalent non-viral sexually transmitted
infection (STI) worldwide, with an estimated 174 million new cases
occurring each year. Approximately five million of these infections occur within the United States [1], and 154 million occur
in resource-limited settings [2]. As many as one-third of female
infections and the majority of male infections are asymptomatic,
often causing trichomoniasis to be considered a self-clearing female
Abbreviations: HE , expected heterozygosity; PCR, polymerase chain reaction;
FISH, fluorescent in situ hybridization; MS, microsatellite; SNP, single nucleotide
polymorphism; SCG, single-copy gene.
夽 Note: Nucleotide sequence data reported in this paper are available
in the EMBL, GenBank and DDBJ databases under the accession numbers:
HM365085–HM365208, DQ321767, DQ321764, XM 001301638, XM 001304496,
XM 001310009, XM 001315268, XM 001317855, XM 001318124, XM 001320341,
XM 001321129, XM 001321414, XM 001321947, XM 001322737, XM 001324024,
XM 001324948, XM 001329311, XM 001329418, XM 001329432, XM 001580544,
XM 001581132, XM 001581222, XM 001582251, and XM 001583217.
∗ Corresponding author at: Department of Medical Parasitology, 341 East 25th
Street, New York, NY 20010, USA. Tel.: +1 212 263 4377, fax: +1 212 263 8116.
E-mail address: [email protected] (J.M. Carlton).
0166-6851/$ – see front matter © 2010 Elsevier B.V. All rights reserved.
doi:10.1016/j.molbiopara.2010.08.006
‘nuisance’ disease [3]. However, trichomoniasis has been associated with a number of significant reproductive health sequelae,
including pelvic inflammatory disease [4] and adverse pregnancy
outcomes, such as premature rupture of membranes, preterm
delivery, and low birth-weight [5,6]. Most importantly, it has been
implicated in increasing sexual transmission of HIV up to twofold
[7,8].
In light of these findings, understanding the transmission
dynamics, strain virulence, and pathology of T. vaginalis infection is critically important for protecting patients – particularly
female patients – from serious disease and threats to reproductive health. However, very little is known about T. vaginalis
genetic diversity or population structure. Antigenic characterization, isoenzyme analysis, repetitive sequence hybridization,
restriction fragment length polymorphism (RFLP), random amplified polymorphic DNA (RAPD), pulsed-field gel electrophoresis, and
sequence polymorphism in ribosomal RNA genes and intergenic
regions have been used to type T. vaginalis isolates in an attempt
to find correlations between genotypes and biologically relevant
phenotypes or geographical distribution [9–16]. Instead of providing clarification, these studies have yielded discordant results for
a number of different phenotypes including metronidazole resistance and geographical distribution [9,10,13,14], the presence of
a linear, double-stranded RNA virus known as TVV [10,13], clin-
M. Conrad et al. / Molecular & Biochemical Parasitology 175 (2011) 30–38
ical manifestation of infection in patients [12,13], and virulence
[9,10,13].
Multilocus genotyping of bacterial and eukaryotic pathogens
has been used successfully to describe population diversity, delimit
species, identify genetic components of important clinical phenotypes, and track the spread of epidemics [17–19]. While RAPD and
RFLP techniques from the pre-genomics era have produced significant amounts of low-cost multilocus data with relative ease,
microsatellite (MS) loci, also known as short tandem repeats, have
been found to be superior tools for most applications [20,21]. These
genetic markers are tandemly repeated 2–6 bp DNA sequences that
display length polymorphism due to changes in the number of
repeat units, and are frequently multi-allelic at each locus. They are
expressed co-dominantly and are abundant throughout eukaryotic
genomes. Crucially, they are generally considered to be neutral alleles that have not evolved under selective pressure, making them
an ideal tool for determining population history. Finally, because
of their high diversity, sensitivity, and multilocus nature, they also
allow for improved accuracy in detection of mixed genotype infections [22]. MS markers have been an important means for studying
population genetics of a number of eukaryotic parasites [17,23–26],
having been used to characterize population structure in Leishmania tropica [27], Trypanosoma cruzi [28], T. brucei gambiense [29],
Plasmodium falciparum [23], P. vivax [30], and Toxoplasma gondii
[26]; to construct a genetic map for Plasmodium falciparum locating regions involved in such phenotypes as drug resistance [31];
and to determine the origins and global dispersal of drug-resistant
parasite strains [32].
Single nucleotide polymorphisms (SNPs) are another type of
genetic marker useful in molecular population genetics. Like MS
markers, they can be highly prevalent and widely distributed
within genomes; in contrast to MS, these markers are usually
bi-allelic due to low mutation rates, which reduce the risk of homoplasy and simplify mutation models. These characteristics make
them especially suited for certain analyses, such as the inference
of phylogenies [33], particularly when they are localized within
single-copy genes (SCG), although careful selection of SCG SNP
markers is important to prevent gene-specific evolutionary history
being mistaken for population evolutionary history. This danger
persists despite the development of sophisticated statistical methods that are robust to a number of confounding influences on
phylogenetic analyses [34]. Therefore, MS and SCG markers must
be validated before they can be adopted as molecular tools for
understanding the diversity and population structure of the parasite under study.
A genome sequence is an ideal resource for anyone intending to
mine genetic loci for use as markers in population genetic studies,
especially of an organism refractory to classical genetics. The draft
T. vaginalis genome, published in 2007 [35], is a ∼160 MB assembly with a core set of ∼60,000 protein-coding genes. Even at 7.2×
coverage, the assembly remains highly fragmented, with ∼74,000
contigs representing the parasite’s six chromosomes. The poor
quality of the assembly is due to such complicating factors as
an apparently rapid, massive expansion of gene families, and the
highly repetitive nature (>65%) of the genome, which rendered
in silico genome assembly challenging. However, this superabundance of repeats suggests a genomic plenitude of MSs that can
be exploited, while also suggesting a comparative lack of singlecopy genes.
Here we report the identification and characterization of 86
microsatellite markers and 21 single-copy genes from the genome
of T. vaginalis, from which a panel of 27 polymorphic genetic markers (21 MS markers and 6 SCG markers) were developed using
seven common laboratory strains of diverse geographical origin,
and six sub-strains. We have adapted our MS typing method to
incorporate a simple and affordable fluorescent labeling protocol,
31
thereby drastically reducing the cost of genotyping and making the
technique readily available to members of the research community. To validate the usefulness of the markers, we have measured
the stability and distribution of the MS, and used phylogenetic and
minimum-spanning network analyses to compare the evolutionary relationships inferred by our panel of markers. Ours is the first
robust series of polymorphic genetic markers for the T. vaginalis
parasite that can be used to classify and monitor lab strains, as well
as provide a means to measure the genetic diversity and population
structure of T. vaginalis field isolates.
2. Materials and methods
2.1. Parasite strains and DNA extraction
A list of the seven geographically widespread T. vaginalis laboratory strains used in this study is shown in Table 1. Two
strains, BRIS/92/HEPU/B7268 and BRIS/92/HEPU/F1623 (abbreviated as B7268 and F1623, respectively) were cultured under drug
selection for a minimum of 255 and 530 days, respectively, and parasites were sampled at various time points during their continuous
culture. DNA extraction for all strains was performed using modification of a published protocol [36]. Approximately 1.5 × 108 cells
from late log phase cultures were harvested at 3000 rpm for 10 min
at room temperature. The cell pellet was resuspended in 0.1×
UNSET lysis buffer (8 M urea, 2% sarkosyl, 0.15 M NaCl, 0.001 M
EDTA, 0.1 M Tris–HCl pH 7.5), and the lysate immediately extracted
twice with an equal volume of phenol:chlorophorm:isoamyl alcohol (25:24:1) and once with an equal volume of chloroform:isoamyl
alcohol (24:1). Nucleic acids were precipitated with 0.6 volumes of
ice-cold isopropanol for 30 min at room temperature or overnight
at 4 ◦ C. Following centrifugation at 12000 rpm for 40 min, the pellet was rinsed three times with 1 mL ice-cold 70% ethanol, dried at
room temperature and resuspended in 50 ␮L low TE buffer, pH 8
(10 mM Tris–HCl, pH 8, 0.1 mM EDTA, pH 8.0) containing 5 ␮g/mL
RNase A.
2.2. Microsatellite marker identification, genotyping and stability
The T. vaginalis genome sequence was mined using version 1.1 of
the genome database resource TrichDB (http://trichdb.org; see Ref.
[37]). The genome was screened for microsatellites (MS) with di-,
tri-, tetra-, penta-, and hexa-nucleotide repeats, using the default
settings of SciRoKo [38] as follows: a fixed penalty of five; allowing
no more than three simultaneous mismatches; requiring a minimum score of 15 and a minimum of three repeats. The MS were
filtered to include only those smaller than 400 bp to ensure that
products would fall within the size standard range during genotyping. Sequences from the regions 200 bp upstream and downstream
of each MS were downloaded and used to design MS-flanking
primers with Primer3Plus [39], using default parameters modified
for 100–450 bp products. Primer sequences were searched against
the complete T. vaginalis genome with Megablast [40] to ensure
uniqueness. A 19 nt tag sequence 5 -GTCGTTTTACAACGTCGTG-3
was added to the 5 end of all forward primers, and the final primer
set was commercially synthesized (Table S1).
All MS markers were amplified as follows: 25 ␮L PCR reactions contained 50 nM forward primer, 1 ␮M reverse primer,
1 ␮M of the primer 5 -GTCGTTTTACAACGTCGTG-3 labeled with
phoshoramidite conjugate HEX or 6-FAM at the 5 end, 1× Mg-free
PCR buffer (Promega, Madison, WI), 0.25 mM dNTPs (New England
BioLabs Ipswich, MA), 1.5 mM MgCl2 , and 0.02 U/␮L GoTaqFlexi
(Promega, Madison, WI). Thermocycler programs were optimized
for each primer combination (Table S1). We tested two different
cycling programs (step and standard) at temperatures ranging from
45 to 60 ◦ C and for each locus selected the program that worked
32
M. Conrad et al. / Molecular & Biochemical Parasitology 175 (2011) 30–38
Table 1
Seven T. vaginalis laboratory strains and six sub-strains used in this study. Several of the commonly used strains are available through the American Type Culture Collection
(ATCC). Six sub-strains isolated during the course of long periods of continuous culture are also shown, with the day of isolation indicated (Day X).
Identifier
Origin
B7RC2
C1:NIH
T1
G3
CI6
BRIS/92/HEPU/F1623 (Day 0)
BRIS/92/HEPU/F1623-Met (Day 350)
BRIS/92/HEPU/F1623-Met (Day 440)
BRIS/92/HEPU/F1623-Met (Day 530)
BRIS/92/HEPU/B7268 (Day 0)
BRIS/92/HEPU/B7268-Met (Day >55)
BRIS/92/HEPU/B7268-Met (Day >55)
BRIS/92/HEPU/B7268-toy (Day >200)
North Carolina, USA
Maryland, USA
Taipei, Taiwan
Kent, UK
Puerto Rico
Brisbane, Australia
Brisbane, Australia
Brisbane, Australia
Brisbane, Australia
Brisbane, Australia
Brisbane, Australia
Brisbane, Australia
Brisbane, Australia
most consistently based on ABI 3130xl sequencer results described
below. Controls included (1) amplification of all primer sets with
commercially available human DNA, (2) DNA prepared from a T.
vaginalis negative vaginal swab to verify primer specificity for the
parasite, and (3) DNA-free PCR reactions. To determine the sensitivity of the genotyping to mixed infections, artificial mixtures of
two strains with genotypes known to differ was evaluated at three
loci (MS70, MS77, and MS135) at 1:1, 2:1 and 3:1 concentrations.
These concentrations allowed for the detection of multiple peaks
in all cases (data not shown). PCR amplicons were run on an ABI
3130xl sequencer with GeneScan-500 LIZ size standard (ABI, Foster
City, CA) for size determination.
2.3. Identification and sequencing of single-copy genes (SCG)
Single-copy genes (SCGs) in the highly repetitive T. vaginalis genome were identified by an all-against-all BLASTP [41]
comparison of protein sequences, excluding those annotated
as hypothetical. Sequences with >60% sequence similarity were
removed from further consideration. The remaining genes were
filtered by size and those ≤3.5 kb retained. Genes were selected
manually from this list by criteria indicating likely polymorphism,
for example, if they were implicated in biologically relevant mechanisms such as virulence. In addition, genes coding for several
proteins identified as being located on the parasite cell surface
[35] were screened for suitability as polymorphic markers. Primers
were designed using Primer3Plus [39] using default parameters
and the final primer set commercially synthesized (Table S2).
All genes were amplified as follows: 50 ␮L PCR reactions contained 1 ␮M forward primer, 1 ␮M reverse primer, 1× Mg-free PCR
buffer (Promega, Madison, WI), 0.25 mM dNTPs (New England BioLabs, Ipswich, MA), 1.5–2 mM MgCl2 , and 0.02 U/␮L GoTaqFlexi
(Promega, Madison, WI). Thermocycler programs (called 2 and
3.5 kb) were optimized for each primer combination using gradient PCRs ranging from 45 to 65 ◦ C (Table S2). Controls included
DNA-free PCR reactions. PCR products were visualized on 1%
agarose gels and those containing single amplicons were purified
using Agencourt AMPure (Beckman Coulter Genomics, Danvers,
MA). The band of interest in PCRs with multiple products were
purified via gel isolation using Wizard SV Gel and PCR Cleanup System (Promega Madison, WI). Sequencing reactions were
performed using BigDye Terminator v 3.1 (ABI, Foster City, CA)
using forward primers, reverse primers, and internal sequencing primers when necessary. All genes were sequenced at ≥2×
coverage. Sequencing reactions were cleaned using Agencourt
CleanSEQ (Beckman Coulter Genomics, Danvers, MA) and analyzed on an ABI 3130xl sequencer. The new sequences from T.
vaginalis lab strains were deposited in GenBank as indicated in
Table S3.
Year
1986
1956
1993
1973
1980s
1992
1992
1992
1992
1992
1992
1992
1992
Availability
References
ATCC #50167
ATCC #30001
Available from J. Carlton
ATCC #PRA-98
BioMed Diagnostics, Inc.
Available from J. Carlton
Available from J. Carlton
Available from J. Carlton
Available from J. Carlton
Available from J. Carlton
Available from J. Carlton
Available from J. Carlton
Available from J. Carlton
[58]
[59,60]
[61]
[62–64]
Unpublished
[65,66]
[65,66]
[65,66]
[65,66]
[66–68]
[66–68]
[66–68]
[66–68]
2.4. Chromosome spreads and fluorescent in situ hybridization
(FISH)
FISH was used to map T. vaginalis MS markers to their chromosomal positions. T. vaginalis cells were prepared using modifications
to a published protocol [42]. Briefly, G3 parasites were grown to
mid-log phase in Diamonds media with 10% horse serum, 1 mM
Colchicine (Sigma, St. Louis, MO) added to 30–50 mL of culture
(∼1.5 × 106 cells/mL), incubated at 37 ◦ C for 6 h, and the cells harvested by centrifugation (all centrifugations were performed at
1000 × g for 5 min at 4 ◦ C). Cells were resuspended in 10 mL of
75 mM potassium chloride, allowed to swell for 5 min at 37 ◦ C,
centrifuged, gently resuspended in 5 mL of fresh Carnoy fixative
(ethanol:chloroform:glacial acetic acid, 6:3:1) at 4 ◦ C for 20 min,
centrifuged again and finally left resuspended in ∼1–2 mL fresh
Carnoy fixative 4 ◦ C over night. Approximately 20 ␮L of the cell suspension were dropped from a height of ∼100 cm onto grease-free
microscopic slides and left to air-dry.
Primers to amplify 1.5–2.5 kb regions spanning each of the MS
loci were designed with Primer3Plus [39] using default parameters
and the final primer set commercially synthesized (Table S3). All
genes were amplified as described for SCGs using either the step
or 2 kb thermocycler program. Amplicons were gel purified using
the Wizard SV Gel and PCR Clean-up System (Promega Madison,
WI), labeled with DIG-11-dUTP using the DIG DNA Labeling Kit
(Roche Applied Science, Indianapolis, IN) and cleaned by ethanol
precipitation.
Freshly prepared T. vaginalis chromosome spreads were briefly
placed in 50% acetic acid solution, dried at 37 ◦ C, incubated in
50 ␮g/mL pepsin (Sigma, St. Louis, MO) in 3 mM acetate buffer with
0.01 M HCl, and washed with PBS pH 7.4 at room temperature for
5 min. The chromosomes were post-fixed with fresh 2% formaldehyde in PBS for 30 min, and incubated in 1% H2 O2 in PBS for 30 min
(all at room temperature). Following dehydration in 3 min washes
of increasing concentrations of methanol (70%, 90%, and 100%) and
subsequent drying, the chromosomes were denatured in 70% formamide in 2× SSC for 5 min at 75 ◦ C under a cover slip and then
immediately dehydrated in a methanol series as described above.
A total of 2 ␮L of each probe (∼20 ng) was denatured in 50 ␮L
of 50% deionized formamide (Sigma, St. Louis, MO) in 2× SSC at
90 ◦ C for 5 min and submerged in ice for 3 min. The probe mixture
was applied to the chromosome slides, a coverslip overlaid and
sealed with rubber cement, and hybridization was left to proceed
overnight in a humid chamber at 37 ◦ C.
High stringency washes were performed at 42 ◦ C in three
changes of 50% formamide (Fluka Buchs, SUI) in 2× SSC for
5 min each, 3× 5 min washes in 2× SSC, and a 5 min wash
in TNT wash buffer (100 mM Tris–HCl, 150 mM NaCl, 0.05%
Tween20, pH 7.5), both at room temperature. At room tempera-
M. Conrad et al. / Molecular & Biochemical Parasitology 175 (2011) 30–38
ture, samples were blocked for 30 min with TNB blocking buffer
(PerkinElmer, Waltham, MA), and then incubated for 60 min in a
humid chamber with anti-digoxigenin antibody conjugated with
horseradish peroxidase (Roche Applied Science, Indianapolis, IN)
diluted 1:1000 in TNB. Slides were washed 3 times for 5 min in
TNT wash buffer at room temperature. The TSA-Plus TMR System
(PerkinElmer, Waltham, MA) was used for tyramide signal amplification according to the manufacturers instructions. Slides were
counterstained with DAPI in Vectashield mounting medium (VectorLabs, Burlingame, CA).
Slides were visualized using a Leica TCS SP2 AOBS confocal
microscope at 63×, and images were merged using LCS Lite v.
2.5 (Leica Microsystems Heidelberg GmbH, Mannheim, Germany).
Chromosomes were identified by their morphological characteristics [42] (see Fig. 1 legend), and chromosome assignment was
performed twice, first with a locus-identifying label, and then a second time in randomized order with labels removed (i.e., blinded).
2.5. Population genetic and evolutionary analyses
GeneMapper 4.0 (ABI, Foster City, CA) was used to score MS allele
sizes. All calls were manually edited to discard data from poorly
amplified reactions and to ensure that proper allele calls were
assigned. Strains were classified as mixtures of two or more genotypes if multiple peaks appeared at two or more loci in duplicate
reactions. HE (expected heterozygosity) was calculated using the
33
n
2
p where p is the frequency of
formula HE = [n/(n − 1)] 1 −
i=1
the ith allele and n is the number of alleles sampled and confirmed
with Arlequin3.11 [43]. A phylogeny using MS markers was inferred
using POPTREE2 [44], a program that constructs phylogenetic trees
from allele frequency data. A neighbor-joining (NJ) tree was constructed using DA [45] distance as a measure of genetic difference
between individuals. Support values for the tree were obtained
by bootstrapping 1000 replicates. A minimum-spanning network
was inferred using Network 4.516 (Fluxus Technology Ltd. 2009)
[46], software developed to reconstruct all possible least complex
phylogenetic trees using a range of data types.
For SNPs, SCG sequence data was aligned to the reference
sequence published in TrichDB and the alignments manually
edited using Sequencher 4.8 (Gene Codes Corporation, Ann Arbor,
MI). Called SNPs were manually verified and included any single nucleotide change that occurred in any single strain. We used
ModelGenerator v. 0.85 [47] to identify appropriate nucleotide substitution models for inference of trees for the six most polymorphic
gene sequences: a set of three surface protein-coding genes TVAG
400860 (GP63a), TVAG 216430 (GP63b), and TVAG 303420 (LLF4),
and a set of three housekeeping genes TVAG 005070 (PMS1), TVAG
302400 (Mlh1a), TVAG 021420 (coronin, CRN), with the number
of gamma categories set at ten. PhyML [48] as part of SeaView v.
4.2.4 [49] was used to infer maximum likelihood (ML) phylogenies reconstructed by applying simultaneous NNI (nearest neighbor
interchange) and SPR (subtree pruning and regrafting) moves on
Fig. 1. Representative images of metaphase-arrested chromosome spreads hybridized with probes specific to microsatellite loci: (top left panel) representative image
showing the T. vaginalis karyotype and chromosome numbering according to a distinct set of karyometrical data [35]; (top right panel) MS20 localized to chromosome
II; (bottom left panel) MS77 localized to chromosome II; (bottom right panel) MS03 localized to chromosome II. The six T. vaginalis chromosomes are morphologically
distinct as follows: chromosome I is the longest and is a subtelocentric/submetacentric type; chromosome II is acrocentric; chromosome III is metacentric; chromosome IV
is metacentric/submetacentric; chromosome V is subtelocentric/acrocentric, although the short arm is not always visible; chromosome VI is the shortest and acrocentric.
34
M. Conrad et al. / Molecular & Biochemical Parasitology 175 (2011) 30–38
five independent random starting trees. Substitution rate categories were set at ten and transition/transversion (Ts/Tv) ratios,
invariable sites and across-site rate variation were selected as indicated by ModelGenerator. Support values for the tree were obtained
by bootstrapping 1000 replicates. Each gene was initially run independently and two manually concatenated DNA sequence data
sets were generated based on their topology. The surface protein
sequences were analyzed using Tamura and Nei’s 1993 nucleotide
substitution model [50] with a Ts/Tv ratio of 2.4, 12 rate categories,
gamma distribution parameter alpha = 0.04, and the proportion of
invariable sites being set at 0.83, while the housekeeping gene
sequences were analyzed with Hasegawa et al.’s 1985 nucleotide
substitution model [51] with a Ts/Tv ratio of 1.91, four rate categories, and the proportion of invariable sites set at 0.97.
3. Results
3.1. Microsatellite identification and development of a simple
genotyping method
We screened the T. vaginalis genome sequence for microsatellites (MS) as an initial step towards developing a panel of markers
for genetic diversity studies of the parasite. In total, 4,630 MSs were
identified with an average length of 30.50 nt ± 1.07 SE, with trinucleotide and pentanucleotide repeats being the most abundant
(37.7% and 25.6%, respectively), followed by tetranuclotide (14.9%),
hexanuclotide (12.3%) and dinucleotide (9.5%) repeats. After further
filtering to exclude MSs present on the same scaffold, we designed
unique primer pairs to 86 MSs for testing (Table S1). Of these 86
MS loci, a total of 21 were found to amplify reliably under our
optimizing conditions and are described below in detail.
We also adapted previously published methods to develop
a simple, reliable and affordable high-throughput MS genotyping protocol for T. vaginalis population genetic studies. First, we
adapted a ‘tagged primer’ technique [52,53] by adding a 19 nt tag
to the 5 end of each forward primer, designed to be recognized by
a fluorescently tagged universal primer. Primers were added in a
1:40:40 ratio (locus-specific forward primer:locus-specific reverse
primer:tagged universal primer) to perform semi-nested PCRs in
which priming was done first with locus-specific reactions, which
were then used as a template for reactions with the labeled primer.
Using one fluorescently tagged universal primer to label all 21
loci greatly reduces genotyping costs. We tested two amplification programs [54] (Table S2) at a range of annealing temperatures,
and validated amplicons by electrophoretic sizing to determine
the most reliable program for each locus. For the latter step HEXlabeled and 6-FAM labeled reactions were multiplexed on the ABI
3130xl, further reducing genotyping costs and increasing the efficiency of the semi-automated, high-throughput methods described
here.
3.2. T. vaginalis microsatellites are stable over many generations
To determine if the microsatellite loci characterized in this study
are stable over long periods (e.g., during continuous culture of the
parasite), we genotyped two T. vaginalis strains that had been maintained in culture for many months. T. vaginalis strain F1623 was
maintained under metronidazole selection for a minimum of 530
days, and strain B7268 was maintained under metronidazole selection for a minimum of 55 days and toyocamycin selection for a
minimum of 200 days (Table 1). We genotyped each strain in duplicate with all 21 MS markers at four separate time points: Day 0,
two Day 55 time points from independent cultures, and Day 255
for strain B7268; at Day 0, Day 350, Day 440, and Day 530 for strain
F1623. No changes were observed at any of the four time points
evaluated (data not shown), indicating that the MS loci are stable
for at least 17 months under highly selective conditions. For all further analyses the data for these different time points were collapsed
and considered as a single isolate.
3.3. Assignment of MS markers to T. vaginalis chromosomes
In order to ensure that our polymorphic MS loci were distributed throughout the T. vaginalis genome, we used fluorescent
in situ chromosome hybridization (FISH) to map MS loci to its six
chromosomes. T. vaginalis chromosome spreads were hybridized
with DIG-labeled locus-specific probes designed to be 1.5–2.5 kb
in length and to span the entire MS locus, and hybridized probes
were visualized using a tyramide signal amplification system. The
chromosomes were identified by their morphological characteristics after two independent analyses of confocal images (Fig. 1).
Although successful signal detection from hybridizations of the
probes was infrequent, most likely due to the single-copy nature
of the loci, 13 of the 21 MS were successfully mapped to single
chromosomes, and the location of a 14th marker (MS06) could
be narrowed down to one of two chromosomes (Table 2). The 14
mapped MS markers are distributed among four of the six chromosomes, with an average of approximately three markers per
chromosome, ranging from two markers on chromosomes I and
IV, to five markers on chromosome II (Table 2). Chromosomes V
and VI, the smallest of the six chromosomes, were not represented
by the mapped markers, although any of the remaining eight MS
markers may be located on them.
3.4. T. vaginalis MS markers from common laboratory strains are
highly polymorphic
We used the 21 MS markers identified above to genotype seven
lab strains of T. vaginalis. Several of the strains are maintained at
the American Type Culture Collection (ATCC) and are commonly
used by T. vaginalis labs around the world. MS loci ranged from
2 to 6 nucleotides in length, and each locus had from 1 to 6 alleles (Table 1), the average being
3.33 alleles
per locus. We used the
n
formula HE = [n/(n − 1)] 1 −
p2 , where p is the frequency
i=1
of the ith allele and n is the number of alleles sampled, to determine heterozygosity of each locus, and we verified this value with
Arlequin 3.11 [43]. This value varied by locus, ranging from 0 to
0.95, with the average HE determined to be 0.66 ± 0.06. One locus
was found to be monomorphic in the lab isolates, but genotyping
of additional clinical isolates (data not shown) revealed it to be
polymorphic. None of the lab strains appeared to be mixtures of
genotypes by our criteria, or in comparison to a simulated mixture
of two lab strains that we used as a control (see Section 2; data
not shown). These results show that MSs in T. vaginalis are highly
polymorphic and can be used as robust markers for genetic studies.
Phylogenetic analyses of the lab isolates using MS data inferred
trees with low to moderate support (Fig. 2a). The basic topology of
the phylogeny correlates well with the topology of the minimumspanning network inferred by Network (Fig. 2b), supporting the
ability of these markers to represent the evolutionary history of
the strains. Topologies from both methods suggest a branch composed of B7RC2 and F1623 that is consistently delineated from the
remaining five isolates. This separation also occurs in the phylogeny
inferred from concatenated sequences of housekeeping genes (see
below).
3.5. Single-copy gene identification and polymorphism
Although MS markers are useful in characterizing genetic diversity due to their highly polymorphic nature, the high mutation
rate associated with them can also be disadvantageous, leading to
M. Conrad et al. / Molecular & Biochemical Parasitology 175 (2011) 30–38
35
Table 2
Characteristics of 21 T. vaginalis MS loci characterized in this study.
Locus
MS01
MS03
MS04
MS06
MS07
MS08b
MS09b
MS10
MS17
MS20b
MS38b
MS44
MS70
MS77
MS94
MS100
MS129
MS135
MS153
MS168
MS184b
Average
Motif
GAATAA
AAAATA
CAA
TTC
ATTAAT
TTC
AGAA
AAAT
TTTTA
TGT
CTT
AT
TA
GA
AT
TG
ATT
TA
AT
TG
TTG
Chromosome
**
I
II*
ND
III or IV*
II*
III*
IV**
IV*
III***
III**
III*
II**
ND
II***
ND
I*
ND
ND
ND
ND
II**
Scaffold
DS113177
DS113181
DS113183
DS113274
DS113206
DS113227
DS113295
DS113349
DS113178
DS113200
DS113502
DS113642
DS114241
DS114317
DS113199
DS113212
DS113270
DS113807
DS113342
DS113392
DS113453
G3 lengtha
Alleles
HE
584,929
437,572
431,961
170,984
244,377
207,319
155,794
128,477
503,923
279,084
91,738
69,902
24,950
22,118
279,741
227,203
172,859
53,461
132,270
115,942
99,852
170
253
250
406
401
270
423
421
206
432
266
247
252
227
209
194
193
248
242
183
254
3
1
3
6
3
3
2
4
4
2
2
4
4
3
2
2
5
4
4
4
5
0.52
0
0.67
0.95
0.76
0.67
0.29
0.87
0.9
0.29
0.29
0.81
0.86
0.67
0.57
0.57
0.86
0.81
0.81
0.81
0.86
211,165
268.7
3.33
0.66
Scaffold size (bp)
ND, not determined.
a
The actual size of MS amplified from reference strain G3 are all within 1–4 bp of the expected size as predicted by in silico analysis of the G3 genome assembly, with the
exception of with the 19 bp tag sequence added to all loci and two markers, MS04 and MS09, which have a 6 and 13 bp difference, respectively.
b
Indicates microsatellite is located within an annotated gene.
*
Localization inferred from one replicate.
**
Localization inferred from two replicates.
***
Localization inferred from ≥3 replicates.
Fig. 2. (A) Neighbor joining tree inferred from microsatellite data using DA distance. Numbers in the tree correspond to non-parametric bootstrap supports (1000 replicates).
(B) Minimum-spanning network inferred from microsatellite data using reduced median setting. (C) Maximum likelihood phylogeny inferred using concatenated surface
protein gene sequences. The log-likelihood of the corresponding phylogenetic model is −5946.1 (D) Maximum likelihood phylogeny inferred using concatenated housekeeping
gene sequences. The log-likelihood of the corresponding phylogenetic model is −7963.0. For both ML phylogenies, the numbers in the trees correspond to non-parametric
bootstrap supports (1000 replicates).
36
M. Conrad et al. / Molecular & Biochemical Parasitology 175 (2011) 30–38
Table 3
SNPs identified in 21 single-copy genes sequenced in seven T. vaginalis strains. Genes whose IDs are underlined were used to generate phylogenetic trees shown in Fig. 2.
Gene ID
Gene name
No. SNPs
Sequence length (bp)
Accession Nos.
TVAG 400860
Clan MA, family M8, leishmanolysin-like
metallopeptidase (GP63a)
Mismatch repair MutL homolog (PMS1)
Mismatch repair MutL homolog (Mlh1A)
Coronin (CRN)
Antigenic protein P1, putative (VSA)
Clan MA, family M8, leishmanolysin-like
metallopeptidase (GP63b)
Vesicular mannose-binding lectin, putative (LLF4)
Vesicular mannose-binding lectin, putative, PS (LLF1)
Vesicular mannose-binding lectin, putative
HIV-1 rev binding protein, putative
Clan CA, family C1, cathepsin L-like cysteine peptidase
Clan CA, family C1, cathepsin L-like cysteine peptidase
Multidrug resistance pump, putative
Tubulin alpha chain, putative
Aspartic peptidase
Tropomyosin isoforms 1/2, putative
Arp2/3, putative
Actin depolymerizing factor, putative
Histone deacetylase complex subunit SAP18, putative
Clan CE, family C48, cysteine peptidase
Conserved hypothetical protein (with PF03388
Domain)
19
1425
HM365121–HM365126, XM 001324948
13
13
7
7a
6
1617
2337
1659
1055
1638
HM365174–HM365178, XM
HM365169–HM365173, XM
HM365115–HM365120, XM
HM365203–HM365208, XM
HM365127–HM365132, XM
5
4
4
4
4
3
3
2
2
2
2
1
1
0
0
1119
1032
1095
750
1233
729
885
834
565
879
801
378
294
640
1119
HM365157–HM365162, XM 001329311
HM365139–HM365144, XM 001329418
HM365151–HM365156, XM 001310009
HM365133–HM365138, XM 001315268
HM365109–HM365114 XM 001321129
HM365103–HM365108, XM 001580544
HM365163–HM365168, XM 001329432
HM365197–HM365202, XM 001322737
HM365179–HM365184, XM 001324024
HM365191–HM365196, XM 001321947
HM365091–HM365096, XM 001318124
HM365085–HM365090, XM 001581222
HM365185–HM365190, XM 001582251
HM365097–HM365102, XM 001583217
HM365145–HM365150, XM 001321414
TVAG
TVAG
TVAG
TVAG
TVAG
005070
302400
021420
364940
216430
TVAG
TVAG
TVAG
TVAG
TVAG
TVAG
TVAG
TVAG
TVAG
TVAG
TVAG
TVAG
TVAG
TVAG
TVAG
303420
291830
086190
171780
485880
228710
291970
184510
459080
414100
087140
192620
166900
094560
309150
a
001301638, DQ321767
00132034, 1DQ321764
001581132
001304496
001317855
All seven SNPs were found within a single isolate that also has a frame-shifting deletion.
homoplasies that may confound true evolutionary histories. Single nucleotide polymorphisms (SNPs), on the other hand, arise less
frequently, are less subject to homoplasy, and with the advent of
high-throughput sequencing methods, are easier to genotype. Since
SNPs located within single-copy genes (SCGs) are most useful for
inferring phylogenies, we mined the T. vaginalis genome for SCGs
using a basic all-versus-all BLASTP of all non-hypothetical proteins, and identified 4,582 genes as suitable candidates. The list
was further reduced to 845 genes by removing all those with a
shared annotated name, and reduced further still on the basis of
size and function. A final experimental set of 21 loci included 18
genes identified through this screening process and three genes
(GP63a, GP63b, and LLF4) included because their putative function as surface proteins suggested that they might be polymorphic.
Amplification and sequencing of all 21 genes from seven of the laboratory strains identified a wealth of SNPs (Table 3). Polymorphism
varied by locus, with some genes highly conserved and exhibiting
no variation (TVAG 094560 and TVAG 309150) and others having
up to 19 SNPs in ∼1.4 kb of sequence (TVAG 400860).
We selected six of the most variable genes (GP63a, PMS1, Mlh1a,
CRN, GP63b, and LLF4) representing housekeeping genes, potential virulence factors and drug targets, for phylogenetic analysis.
(Although TVAG 364940 [VSA] was found to be more polymorphic
than TVAG 216430 [GP63b] and TVAG 303420 [LLF4], the polymorphisms were restricted to a single isolate with a frame-shifting
deletion, and so we excluded this gene.) After determining appropriate nucleotide model parameters, we used PhyML [48] to infer
the phylogeny of each gene separately. The three surface protein
genes (GP63a, GP63b, and LLF4) had similar topologies, as did the
three housekeeping genes (PMS1, Mlh1a, and CRN), although these
topologies differed between the two groups. Using the similarity
between the topologies, we arranged the sequence data into two
concatenated data sets, one composed of the three putative surface
protein-coding genes (GP63a, GP63b, and LLF4), and one composed
of the three housekeeping genes (PMS1, Mlh1a, and CRN). The surface protein data set was composed of 4182 unambiguously aligned
positions with 30 polymorphic sites, translating to 24 variable sites
in the protein sequence, one of which has three amino acid variants,
while the housekeeping data set was composed of 5619 unambiguously aligned positions, including 34 polymorphic positions,
translating to 23 variable sites in the protein sequence. The two
data sets were analyzed using Tamura and Nei’s [50] and Hasegawa
et al. [51] nucleotide substitution models, respectively. The two
trees share little topology (Fig. 2c and d), probably as a result of
the different selective pressures and evolutionary forces the two
sets of genes experience. The phylogeny inferred from the concatenated housekeeping sequence has better bootstrap support than
that of the concatenated surface protein sequence; in addition, it
more closely resembles the phylogenetic and minimum-spanning
network topologies inferred from microsatellite data (Fig. 2a, b,
and d), suggesting that the separation of B7RC2 and F1623 from
the remaining strains is the best representation of their ancestral history under assumed neutral selection pressure. Including
additional lab strains or isolates would likely clarify relationships
between individual strains that are currently ambiguous.
4. Discussion
A draft of the first T. vaginalis genome sequence was published
in 2007 [35], opening the way for development of new genetic tools
to understand the biology of this neglected pathogen. Although
Sanger sequencing coverage of the genome is high at ∼7.2×, the
presence of hundreds of gene families and thousands of repetitive
elements that show extreme similarity to each other (average pairwise difference ∼2.5%) hinder the generation of a more complete
assembly of the genome. As a result, identifying the single-copy
genes and unique MS markers reported in this study has been a
challenging and non-trivial exercise.
Here we have described a novel suite of 27 robust, reproducible,
and inexpensive genetic markers for investigating the population
genetics of T. vaginalis, including the first reported MS markers and
a panel of diverse single-copy genes. In addition, we have mapped
the MS loci to individual chromosomes, verified their stability, and
performed minimum-spanning network and phylogenetic analysis to validate their usefulness for inferring the evolutionary
relationship of strains. We have similarly identified polymorphic
single-copy genes and also demonstrated their use in phylogenetic
studies.
Our results show that the 21 MS loci that we identified and
successfully optimized for genotyping in seven laboratory strains
M. Conrad et al. / Molecular & Biochemical Parasitology 175 (2011) 30–38
and six sub-strains exhibit a high degree of diversity, and are distributed among at least four of the six chromosomes comprising
the T. vaginalis genome. We have shown these markers to be stable
in continuous culture over extended periods of time while under
drug selection, suggesting that the mutation rate at these loci is
slow enough to prevent extensive homoplasy, and supporting their
usefulness as markers.
By identifying single-copy genes coding for protein sequences
lacking strong similarity to other protein sequences encoded by the
genome, we have been able to eliminate complications arising from
the use of paralogs (genes related by duplication within a genome),
ensuring that phylogenies represent locus-specific evolutionary
history and therefore better reflect the ancestral relationship of
T. vaginalis isolates. We have also selected genes with diverse
functions, including housekeeping genes and putative surface proteins, thereby representing the history of loci evolving under a
range of selective pressures. Combining sequence data from these
genes allows us to accurately represent the evolutionary history
of the genome as whole, rather than of select loci. In addition, we
have identified genes with sufficient polymorphism to distinguish
between closely related strains, making these markers useful for
intra-specific, population-level analyses.
The six genes that we have selected for phylogenetic analysis demonstrate different evolutionary relationships among the
strains. The housekeeping gene phylogenies strongly support
strains B7RC2 and F1623 being more closely related to each other
than to the other isolates, a relationship that is maintained in phylogenies of the individual genes that make up the concatenated
data set. This association is also found in ancestral relationships
inferred from microsatellite data using two different methods of
analysis. In contrast, the surface genes indicate a closer relationship between B7268 and F1623. This relationship is only apparent
in the GP63a gene, as the other individual genes have low support
values at all nodes (data not shown). This may indicate that the
relationship of these genes is more influenced by selective pressure than common ancestry. Pressure for diversification of surface
proteins to adapt to host immune responses and/or to be able to
colonize a host may overshadow other aspects of their evolutionary history, making their ancestral relationship ambiguous in this
small data set. Evaluating additional clinical samples may better
explain the lack of overlap in tree topologies and provide a clearer
picture of the forces driving the relationships between the genes of
different isolates.
During our MS marker development, we employed a costsaving method that utilized one universal tagged primer to label
PCR amplicons of the MS loci [52,53,55], making the technique
accessible to more researchers, particularly those in resourcelimited settings. The availability of these reproducible markers will
allow comparison and merging of data sets from different research
projects, enabling a more comprehensive investigation of the population diversity of T. vaginalis through better representation of
widespread geographical locations, population demographics, and
overall parasite diversity.
As a result of this study and future parallel population-level
diversity studies, those investigating T. vaginalis will be able to
examine the extant population structure and diversity of the parasite to better select strains when studying specific biological
functions. Similarly, this genotyping method will be invaluable for
resolving conflicting experimental results by helping to standardize the influence of genetic background on variation in quantitative
traits. Another powerful use of these markers will be to elucidate
the role of meiosis in the T. vaginalis lifecycle [54] by providing a
means to detect recombinants [56] and to utilize population genetic
models to detect other evidence for or against recombination, such
as linkage disequilibrium [23] or the detection of haplotypes in
significantly excessive frequency [57].
37
T. vaginalis remains a ‘neglected’ pathogen in part due to an
incomplete understanding of the role it has played in pathology
and in facilitating co-infection and transmission of diseases that
are of great public health concern (e.g., HIV/AIDS). As a result, there
is a paucity of tools for elucidating its mechanisms of colonization,
pathogenesis, and drug resistance. The development of this panel of
genetic markers is a step towards rectifying this neglect and raising
awareness of the complexity of the parasite’s genetics, leading to
better explanation of the wide range of severity of symptoms and
manifestations of pathology.
Acknowledgements
We thank Drs. Shehre-Banoo Malik for assistance in phylogenetic analyses, Patrick Sutton for valuable input in the writing of
the manuscript, and Ute Frevert for assistance with the microscopy.
We would also like to recognize the Confocal Microscopy Facility of
the Department of Medical Parasitology for the use of the Leica TCS
SP2 AAOBS confocal microscope. M.C. was partially funded under a
National Institutes of Health training grant T32AI007180-27. This
study was funded by a National Institutes of Health award to J.M.C.
under the Advancing Novel Science in Women’s Health Research
program announcement, grant number R21 AI083954-01. J.T. and
Z.Z. were supported by the Czech Ministry of Education (LC07032,
MSM 0021620858).
Appendix A. Supplementary data
Supplementary data associated with this article can be found, in
the online version, at doi:10.1016/j.molbiopara.2010.08.006.
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Molecular & Biochemical Parasitology 176 (2011) 135–137
Contents lists available at ScienceDirect
Molecular & Biochemical Parasitology
Short technical report
Fluorescence in situ hybridization (FISH) mapping of single copy genes on
Trichomonas vaginalis chromosomes
Zuzana Zubáčová a , Vladimír Krylov b , Jan Tachezy a,∗
a
b
Charles University in Prague, Faculty of Science, Department of Parasitology, Vinicna 7, 128 44 Prague, Czech Republic
Charles University in Prague, Faculty of Science, Department of Cell Biology, Vinicna 7, 128 44 Prague, Czech Republic
a r t i c l e
i n f o
Article history:
Received 19 October 2010
Received in revised form
20 December 2010
Accepted 21 December 2010
Available online 30 December 2010
Keywords:
Trichomonas vaginalis
Chromosome
FISH
a b s t r a c t
The highly repetitive nature of the Trichomonas vaginalis genome and massive expansion of various
gene families has caused difficulties in genome assembly and has hampered genome mapping. Here,
we adapted fluorescence in situ hybridization (FISH) for T. vaginalis, which is sensitive enough to detect
single copy genes on metaphase chromosomes. Sensitivity of conventional FISH, which did not allow
single copy gene detection in T. vaginalis, was increased by means of tyramide signal amplification. Two
selected single copy genes, coding for serine palmitoyltransferase and tryptophanase, were mapped to
chromosome I and II, respectively, and thus could be used as chromosome markers. This established protocol provides an amenable tool for the physical mapping of the T. vaginalis genome and other essential
applications, such as development of genetic markers for T. vaginalis genotyping.
© 2011 Elsevier B.V. All rights reserved.
Trichomonas vaginalis is a flagellated protist belonging to the
Parabasala group. It is an obligate human parasite that causes the
most prevalent non-viral, sexually transmitted infection of the urogenital tract [1]. The parasite reproduces asexually by binary fission,
although the presence of sexual processes in T. vaginalis has also
been suggested [2,3]. The mitosis of T. vaginalis is closed, as the
nuclear envelope remains intact with the formation of an extranuclear spindle [4]. During mitosis, nuclear chromatin condenses into
six chromosomes that can be distinguished according to their size
and morphology [2]. Metaphase chromosomes consist of a pair
of parallel sister chromatids; each chromatid bears its own constriction that is interpreted as a centromere [2]. This chromatid
arrangement is considerably different from the typical monocentric
chromosomes of higher eukaryotes.
Sequencing of the T. vaginalis genome allows the estimation of
its size at about 160 Mb, with a core set of about 60,000 genes, representing the highest coding capacities in eukaryotes [5]. However,
the highly repetitive character of the genome, including 59 repeat
families that constitute almost 39 Mb of the genome, and massive
expansion of various gene families caused difficulties in genome
assembly and hampered gene mapping. Therefore, applications of
genetic techniques such as fluorescence in situ hybridization (FISH)
are essential to improve correct joining of contigs and allow their
Abbreviations: FISH, fluorescence in situ hybridization; TSA, tyramide signal
amplification.
∗ Corresponding author. Tel.: +420 221951811.
E-mail address: [email protected] (J. Tachezy).
0166-6851/$ – see front matter © 2011 Elsevier B.V. All rights reserved.
doi:10.1016/j.molbiopara.2010.12.011
mapping on T. vaginalis chromosomes. Moreover, the physical location of single copy genes provides a tool for T. vaginalis genotyping
with applications in evolutionary studies, comparative genomics
and clinical praxis.
Previously, FISH has been employed in T. vaginalis for the mapping of multicopy genes coding for rRNA that are present in its
genome in hundreds of copies [5,6]. These genes are clustered to a
single chromosome IV, which facilitates their easy detection [5].
However, our attempts to detect single copy genes on T. vaginalis chromosomes using conventional FISH were unsuccessful.
In this report, we describe a protocol for highly sensitive FISH,
adapted for the T. vaginalis genome, which allows for the visualization of single-copy genes on metaphase chromosomes of this
parasite. The sensitivity of the system was increased by coupling
FISH with horseradish peroxidase-catalyzed tyramide signal amplification (FISH-TSA).
To establish a FISH-TSA technique for trichomonad chromosomes, we selected four nucleotide sequences with similar
GC-content (42–49%) as probes, coding for asparaginase-like
threonine peptidase (TVAG 258340), acetylornithine aminotransferase, putative (TVAG 258770), serine palmitoyltransferase
(TVAG 388650), and tryptophanase (TVAG 054490). The presence
of a single gene copy for each selected sequence in T. vaginalis
genome was verified by BLAST searches at Tricho DB database
(http://trichdb.org/trichdb/). As a positive control, we used the 28S
rRNA gene (AF202181), which is present in 250 copies. Unlabeled
probes were obtained by PCR amplification using trichomonad
genomic DNA as a template. The DNA was isolated from the T1
strain (kindly provided by J.-H. Tai, Institute of Biomedical Sci-
136
Z. Zubáčová et al. / Molecular & Biochemical Parasitology 176 (2011) 135–137
Fig. 1. Mapping of single-copy genes in the Trichomonas vaginalis genome by FISH-TSA. (A) Metaphase spread with red signals corresponding to a 1419 bp probe hybridized to
serine palmitoyltransferase gene on chromosome I. (B) Visualization of the serine palmitoyltransferase gene in an interphase nucleus. (C) The result of hybridizing a 1377 bp
probe to the tryptophanase gene on metaphase chromosome II. (D) Mapping of a multicopy gene for the 28S rRNA was conducted as a positive control. The probe hybridized
to chromosome IV.
ences, Taipei, Taiwan) with the High Pure PCR Template Preparation
kit (Roche). The primers for gene amplification were designed to
obtain complete sequences of 921–1419 bp (single copy genes)
and 1574 bp (28S rRNA gene). The PCR products were purified
from the gel using QIAquick Gel Extraction Kit (Qiagen), labeled
by digoxigenin-11-dUTP (Roche) using DecaLabel DNA Labeling Kit
(Fermentas), and purified using QIAquick Gel Extraction Kit (Qiagen) in a final volume of 50 ␮l.
The metaphase chromosomes from colchicine-treated T. vaginalis cells were obtained using modifications to the described
spreading technique [7]. T. vaginalis T1 cells were grown in Diamonds TYM media supplemented with 10% horse serum. The
trichomonads in logarithmic growth phase (about 3 × 107 cells
in 20 ml) were then incubated in the same media with 1 mM
colchicine (Sigma) for 6 h at 37 ◦ C and harvested by centrifugation. Cells were hypotonised in 10 ml of 75 mM potassium chloride
for 5 min at 37 ◦ C, spun down, resuspended in 5 ml of Carnoy fixative (ethanol–chloroform–acetic acid, 6:3:1) at 4 ◦ C for 20 min,
spun down, resuspended in 1–2 ml Carnoy fixative and left at 4 ◦ C
overnight. The cell suspension (20 ␮l) was then dropped from a
height of about 20 cm onto microscope slides (Star Frost, Waldemar Knittel Glasbearbeifungs GmbH, Braunschweig, Germany) and
left to dry.
The chromosome spreads were briefly submerged in 50% acetic
acid to remove the cytoplasmic residues, dried, incubated for 5 min
at 37 ◦ C in 50 ␮g/ml pepsin (Sigma) in 3 mM potassium acetate
and 0.01 M HCl to excise proteins, and then washed for 5 min with
phosphate-buffered saline pH 7.4 (PBS) at room temperature. The
chromosome spreads were post-fixed with fresh 2% formaldehyde
in PBS for 30 min at room temperature (after this step, the spreads
could be stored at −20 ◦ C for one month for later use). To remove
any endogenous peroxidase activity, the spreads were incubated
in 1% H2 O2 in PBS for 30 min at room temperature, dehydrated in
series of 70%, 90% and 100% methanol, and air-dried. Subsequently,
the chromosomes were denatured in 50 ␮l of 70% deionized formamide (Sigma) in 2× SSC for 5 min at 75 ◦ C under a coverslip
and dehydrated again in methanol, as described above. Each probe
(20 ng in 2 ␮l) was denatured in 50 ␮l of 50% formamide in 2× SSC
at 90 ◦ C for 5 min and immediately cooled down in ice. Denatured
probe was applied on the chromosome spread, with a coverslip, and
sealed with rubber cement. Hybridization proceeded overnight in
a humid chamber at 37 ◦ C.
The chromosome spreads were washed at high stringency 3
times for 5 min at 42 ◦ C using 50% formamide (Fluka) in 2× SSC, following 3 × 5 min washes in 2× SSC at room temperature, and 5 min
in TNT buffer (100 mM Tris–HCl, 150 mM NaCl, 0.05% Tween-20, pH
Z. Zubáčová et al. / Molecular & Biochemical Parasitology 176 (2011) 135–137
7.5). The spreads were blocked for 30 min in TNB blocking buffer
(PerkinElmer) and incubated for 60 min with anti-digoxigenin
antibody conjugated with horseradish peroxidase (Roche) diluted
1:1000 in TNB buffer at room temperature. Slides were washed
3 × 5 min in TNT buffer at room temperature. The tyramide signal amplification was performed using the TSA – Plus TMR System
(PerkinElmer), according to the manufacturer’s manual. Slides were
counterstained with DAPI in Vectashield mounting medium (VectorLabs). Chromosomes were observed using an IX81 microscope
(Olympus) equipped with an IX2-UCB camera. Images were processed using Cell® software (Olympus).
The high sensitivity of FISH-TSA allows the successful visualization of single-copy genes in a Trichomonas vaginalis karyotype
with minimal background. The gene coding for serine palmitoyltransferase was mapped to the longest chromosome, chromosome
I (subtelocentric/submetacentric type). The specific signal was
detected on the longer chromatid arm, close to the constriction
of each chromatid (Fig. 1A). The signal was frequently observed
only on one of the two sister chromatids (about 20% of total
spreads). When paired doublets were observed (about 5% spreads),
a signal was usually asymmetric, with lower intensity on one of
the chromatids. The strong signal for serine palmitoyltransferase
was also detected on interphase nuclei (Fig. 1B). The single-copy
gene tryptophanase was identified on acrocentric chromosome II
(Fig. 1C). The signal was observed close to the center of sister
chromatids of this chromosome, which do not possess apparent
constriction. As in the case of serine palmitoyltransferase, the signal was often observed only on one of the sister chromatids. Unlike
single-copy genes, the probe against multiple 25S rRNA genes
frequently labeled both sister chromatids (85% of total spreads).
The signal corresponding to 25S rRNA genes was observed as a
single spot on chromosome IV, and it was associated with chromatid constriction (Fig. 1D), as observed previously for 18S rRNA
genes [5].
The adaptation of FISH-TSA provided substantial improvement in sensitivity over conventional FISH for the detection of
genes in the T. vaginalis genome. However, the signal was limited by the length of the probe, as has been observed for other
cell lines [8,9]. Probes for both successfully mapped genes were
over 1300 bp in length (1377 bp, tryptophanase; 1419 bp, serine
palmitoyltransferase gene), whereas two shorter probes against
asparaginase-like threonine peptidase (921 bp) and acetylornithine
aminotransferase (1203 bp) repeatedly did not detect the corresponding genes (data not shown). When compared to plant and
human genomes, for which probes smaller than 1 kb have been successfully used [8], observed sensitivity of FISH-TSA in T. vaginalis
is somehow lower. Beside a GC content, which was comparable
for T. vaginalis (42–49%) and human (41–49%) probes [8], the most
important factor that influences FISH sensitivity is chromatin compactness [9]. Thus, to estimate the level of chromatin condensation
in T. vaginalis metaphase chromosomes, we calculated the ratio
between T. vaginalis genome size (160 Mbp) and total length of
metaphase chromosomes (9.4 ␮m), which is about 17 Mbp ␮m−1 .
This value is comparable or lower than the ratio values known for
human (26.6 Mbp ␮m−1 ) and plants (40.6–249.6 Mbp ␮m−1 ) [9].
Therefore, the observed limitation of probe length in T. vaginalis
137
cannot simply be explained by genome compactness; other factors,
such as the specific character of cellular and nuclear membranes
and cytoplasm that cover the chromosomes on spreads and the specific structure of chromatin related to strongly divergent histones
[10], might affect the availability of the targeted genes for the probe
hybridization. The chromosomes of a number of parasitic protists,
including related intestinal parasite Giardia intestinalis, could be
routinely separated by pulse field gel electrophoresis (PFGE) [11].
Notably, although separation of six T. vaginalis chromosomes have
been reported [12], estimated sizes of six separated DNA bands
(in total 16.2 Mbp) did not correspond to the trichomonad genome
size. Our attempts, and the attempts of other laboratories [13], did
not lead to separation of genomic DNAs corresponding to chromosomes. Considering the large size of T. vaginalis genome (160 Mb),
it is likely that chromosomes of T. vaginalis are too large to be
separated by PFGE that is limited by about 10 Mbp.
In conclusion, we adapted a highly sensitive FISH-TSA method
for the detection of single-copy genes on metaphase T. vaginalis
chromosomes. This method provides an amenable tool for the
joining of available genomic contigs, the physical mapping of
single-copy genes and the definition of physical linkage groups, the
study of genome rearrangements, and the genotyping of T. vaginalis
strains.
Acknowledgement
This study was supported by the Czech Ministry of Education
(LC07032, MSM0021620858) to J.T.
References
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1997;33:131–5.
[3] Malik SB, Pightling AW, Stefaniak LM, et al. An expanded inventory of conserved
meiotic genes provides evidence for sex in Trichomonas vaginalis. PLoS One
2008;3:e2879.
[4] Brugerolle G. Etude de la cryptopleuromitose et de la morphogenese de division
chez Trichomonas vaginalis et chez plusieurs de trichomonadines primirives.
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[5] Carlton JM, Hirt RP, Silva JC, et al. Draft genome sequence of the sexually transmitted pathogen Trichomonas vaginalis. Science 2007;315:207–12.
[6] Torres-Machorro AL, Hernández R, Alderete JF, López-Villaseñor I. Comparative
analyses among the Trichomonas vaginalis, Trichomonas tenax, and Tritrichomonas foetus 5S ribosomal RNA genes. Curr Genet 2009;55:199–210.
[7] Zubacova Z, Cimburek Z, Tachezy J. Comparative analysis of trichomonad
genome sizes and karyotypes. Mol Biochem Parasitol 2008;161:49–54.
[8] Schriml LM, Padilla-Nash HM, Coleman A, et al. Tyramide signal amplification (TSA)-FISH applied to mapping PCR-labeled probes less than 1 kb in size.
Biotechniques 1999;27:608–13.
[9] Khrustaleva LI, Kik C. Localization of single-copy T-DNA insertion in transgenic shallots (Allium cepa) by using ultra-sensitive FISH with tyramide signal
amplification. Plant J 2001;25:699–707.
[10] Marinets A, Müller M, Johnson PJ, et al. The sequence and organization of the
core histone H3 and H4 genes in the early branching amitochondriate protist
Trichomonas vaginalis. J Mol Evol 1996;43:563–71.
[11] Upcroft JA, Krauer KG, Upcroft P. Chromosome sequence maps of the Giardia
lamblia assemblage A isolate WB. Trends Parasitol 2010;26:484–91.
[12] Lehker MW, Alderete JF. Resolution of six chromosomes of Trichomonas
vaginalis and conservation of size and number among isolates. J Parasitol
1999;85:976–9.
[13] Upcroft JA, Delgadillo-Correa MG, Dunne RL, et al. Genotyping Trichomonas
vaginalis. Int J Parasitol 2006;36:821–8.
5. Unpublished data
Zubacova, Z. and J. Tachezy (2011). Histone H3 variants in Trichomonas vaginalis.
41
Histone H3 variants in Trichomonas vaginalis
Histones are DNA - binding proteins that evolved from archaebacterial ancestors and
function in eukaryotic genome packaging. Octameres composed of core histones H2A, H2B,
H3 and H4 form basic units of chromatin called nucleosomes (Dalal et al. 2007;Henikoff and
Ahmad 2005). Although histones are remarkably conserved proteins, several variants of H2A
and H3 have arised. Histone variants differ from core histones in their protein sequences,
distinct localization in chromatin and specialized functions. Histone H2A can be replaced by
H2AZ destabilizing nucleosome structure or H2AX playing role in DNA repair and T cells
differentiation (Redon et al. 2002).
Histone H3 can be replaced by H3.3 or cenH3 that are deposited in active genes and
centromeres, respectively. Histone H3.3 has similar protein sequence as core H3. It is
characterized by lysine methylation and substitues core H3 in nucleosomes in
transcriptionally active chromatin. Recent experiments showed that H3.3 containing
nucleosomes occur also in pericentric heterochromatin and telomeres but its role here is
unclear (Szenker et al. 2011; Talbert and Henikoff 2010). On the other hand, cenH3 is highly
diverged from core H3 especially in N-terminal part of the protein and shares 50-60%
sequence identity only within C-terminal histone fold domain. CenH3 plays crucial role in
chromosome segregation. It is an epigenetic marker of centromeres, a structure that is
defining feature of eukaryotic chromosomes (Ahmad and Henikoff 2002). CenH3
nucleosomes form basement for kinetochore complex that assembles on centromeres and
mediates capture of chromosomes by spindle microtubules and their migration to poles during
cell division. CenH3s were identified in all eukaryotes studied to date with the exeption of
parasite Trypanosoma brucei (Lowell and Cross 2004). Further specific variants of histone H3
have evolved in parasitic protists Trypanosoma brucei and Giardia intestinalis. T. brucei
histone H3V with unknown function is enriched at telomeres (Lowell and Cross 2004). G.
intestinalis histone H3B probably marks noncentromeric heterochromatin ((Dawson et al.
2007).
Trichomonas vaginalis is a human parasite from Parabasala group of protists. Its
repetitive 160 Mb genome is divided into six chromosomes (Carlton et al. 2007). Genes for
all four core histones H2A, H2B, H3, H4 together with three H3 variants can be found in T.
vaginalis genome database by product name and BLAST search (http://trichdb.org/trichdb/).
Per haploid genome, there are 17 copies of histone H2A, 14 copies of H2B, 21 copies of H4
and 23 intronless genes coding for putative histone H3. Twenty trichomonad histones H3 are
identical in their protein sequences and three variants are different (Figure 1a.). Selected H3
variants were aligned with homologs from other organisms using ClustalX ((Thompson et al.
1997) (Figure 1b.).
We examined protein sequences and nuclear localization of three T. vaginalis H3
variants to find centromeric marker cenH3. T. vaginalis strain Tv T1 (kindly provided by J.H. Tai, Institute of Biomedical Sciences, Taipei, Taiwan) was used in the study. Cells were
grown in TYM medium with 10% heat – inactivated horse serum (pH 6, 2) at 37°C (Diamond
1957). We studied experimentally chromatin localization of the following T. vaginalis
histones H3:

TVAG_270080 as core histone H3 according to the alignment (139 amino acids in
lenght) which was used as a control.

TVAG_087830 as H3 variant that differs from core H3 in six amino acid residues
(139 amino acids in lenght).

TVAG_224460 as divergent H3 variant (155 amino acids in lenght). It has N-terminal
extension when aligned to core H3 and nonconserved N-terminal tail.

TVAG_185390 as second the divergent variant that can not be aligned to core histone
H3 at the N-terminus (135 amino acids in lenght). It does not have N-terminal
extension (Figure 1a.).
Depositions of H3 variants in T. vaginalis chromatin were elucidated using their tagged
versions. Genes were PCR amplified, cloned and expressed in trichomonads. The template
genomic DNA was isolated from trichomonads using High Pure PCR Template Preparation
kit (Roche). PCR products were purified from the gel using QIAquick Gel Extraction Kit
(Qiagen). Amplified genes were introduced into TagVag vector with a hemagglutinin (HA)
tag at the 3´ end. Trichomonads were electroporated with histone H3_HA constructs and
maintained as described in Sutak et al. (Sutak et al. 2004). After confirmation of expression of
HA – tagged histones of expected size of 20 kDa in transfected trichomonads by Western
blotting (data not shown), the localization of H3 variants was examined by indirect
immunofluorescence using monoclonal antibody against HA - tag. Culture of trichomonads
enriched in mitotic cells was obtained according to the protocol described in Torres-Machorro
et al. (Torres-Machorro et al. 2009). Metaphase chromosomes were obtained by cultivation of
trichomonads with 1mM colchicine for 6 hours followed by 5 minutes of hypothonic swelling
in 75mM KCl (Zubacova et al. 2008). Microscopic preparations were prepared according to
Sutak et al. (Sutak et al. 2004). Cells were placed on glass slides coated with 3aminopropyltriethoxysilane (Sigma) and fixed with methanol (5 min) and acetone (5 min) at –
20°C. Cells were blocked for 1 hour in PBS/0.25% BSA (Sigma)/0.25% cold water skin
gelatin (Sigma)/0.05% Tween 20 (Sigma) and then incubated for 1 hour or alternatively
overnight with mouse anti-HA monoclonal antibody diluted in blocking buffer to stain
overexpressed histones H3. In double-labeling experiments, rabbit anti-monomethylated
H3K4 antibody (Millipore) was used. After PBS washing, secondary donkey anti-mouse
Alexa Fluor-488 (green) and donkey anti-rabbit Alexa Fluor-546 (red) antibodies (Molecular
Probes) were added for 1 hour. After washing in PBS, the cells were mounted in VectaShield
mounting medium with DAPI (VectorLabs). Fluorescence microscopy was performed using
IX81 microscope, photographed with an IX2-UCB camera and processed using Cell^R
software (Olympus). Cell section images were analysed using ImageJ (NIH, Bethesda, MD,
USA) program to determine the distribution of histone H3 variants.
Despite histones belong to the most conserved proteins, former study revealed
remarkably greater sequence diversity of core H3 and H4 from protists when aligned to their
counterparts from multicellular organisms (Marinets et al. 1996). The highest sequence
identidy of trichomonad histone H3 TVAG_270080 with histones from higher eukaryotes was
82% thus we conclude that this histone is core H3 homolog. On microscopy preparations, it
was distributed homogenously throughout the whole interphase nuclei as well as whole
metaphase chromosomes (Figures 2. and 3.).
The protein sequence of H3 variant TVAG_087830 is almost identical with that of
core H3 except six amino acids substitutions that is typical for histone variant H3.3, a marker
of transcriptionally active chromatin (Figure 1a.) (Ahmad and Henikoff 2002; Henikoff and
Ahmad 2005). This H3 variant was localized in trichomonad chromatin as many small foci
(Figure 2.). Activation of transcription is indicated by histone lysine methylation (Martin and
Zhang 2005). To confirm experimentally that histone TVAG_087830 defines actively
transcribed regions we used anti-monomethyl H3K4 antibody immunostaining to label active
chromatin in nuclei of T. vaginalis. Anti-monomethyl H3K4 stained discrete foci that colocalized with HA-tagged TVAG_087830 H3 variant (Figure 4.).
Centromeric H3 variant, cenH3, is the only stable epigenetic marker of centromeres,
structures that characterize eukaryotic chromosomes and have essential role in genome
segregation during cell division (Ahmad and Henikoff 2002). With the exception of
Trypanosoma brucei, all eukaryotes studied to date have cenH3. In addition, the role of
telomeric T. brucei histone H3V as a cenH3 was not completely exluded (Lowell and Cross
2004). Although, centromeric H3 variants lack conserved sequence motif, based on several
protein sequence criteria that chacterize majority of cenH3s (Ahmad and Henikoff 2002;
Henikoff and Ahmad 2005) we considered the posibility that one of the two divergent histone
H3 variants TVAG_224460 or TVAG_185390 might be marker of trichomonad centromeres.
i.) All studied eukaryotes have only one gene copy in their genomes coding for cenH3
(Cervantes et al. 2006). Both trichomonad cenH3 candidates are single copy genes. ii.)
CenH3s have divergent N-terminus with typical extension when aligned with canonical
histone H3 (Dawson et al. 2007). Both trichomonad putative cenH3s have divergent Nterminal part of the protein but only TVAG_224460 variant has N-terminal extension of ten
aminoacids. Variant TVAG_185390 is shorter of four amino acids residues than core H3
(Figure 1a.). iii.) CenH3s typically share only 50-60% identity within C-terminal histone fold
domain of core H3 (Cervantes et al. 2006; Lowell and Cross 2004). T. vaginalis putative
cenH3s share 60% and 61% sequence identity with core H3 within HFD. iv.) All
experimentally validated centromeric H3 variants have at least one amino acid residue
insertion in loop 1 of secondary protein structure. One amino acid residue insertion is present
in loop 1 of variant TVAG_185390. Variant TVAG_224460 has no amino acid insertion in
loop 1. v.) Conserved glutamine residue in α-1 helix of core histone H3 is often absent in
cenH3s (Lowell and Cross 2004). This glutamine is absent in TVAG_185390 but not in
TVAG_224460 variant (Figure 1b). Although both proteins display some characteristics of
cenH3, their immunofluorescence pattern was completely different. Surprisingly,
immunostaining of H3 variant TVAG_185390 showed the localization along the entire
chromosomal arms and in whole interphase nuclei like core H3. This localization does not
correspond to cenH3 (Figures 2. and 3.).
Second cenH3 candidate TVAG_224460 was localized to six peripheral dots what
corresponds to six chromosomes in G1 phase of the mitosis. Trichomonad cenH3 was
localized as peripheral double dots in G2 phase (with duplicated chromatin). The same
fluorescence pattern of cenH3 was observed previously during Tetrahymena termophila
mitosis (Cervantes et al. 2006). We concluded that this H3 variant marks sites of the
centromeres in T. vaginalis chromatin (Figure 2.).
Our results give support to former observation that trichomonad chromosomes are
monocentric rather than holocentric with diffuse kinetochores (Drmota and Kral 1997).
Immunofluorescence pattern of cenH3 staining differs remarkably between monocentrics and
holocentrics. CenH3 staining of monocentric chromosomes looks like distinct foci
corresponding to the number of chromosomes, whereas cenH3 staining in holocentric
chromosomes is localized as a band on the edge of each sister chromatid facing towards
spindle microtubules (Dernburg 2001). Such pattern we have not seen in T. vaginalis.
Furthermore, in monocentric chromosomes centromeres are visible as a primary constrictions
that are absent in holocentrics (Ahmad and Henikoff 2002; Guerra et al. 2010; Maddox et al.
2004). Secondary constrictions are seen on chromosomes in addition to primary constrictions.
In mitotic metaphase T. vaginalis chromatin condenses into six chromosomes and four of
them bear clearly visible constriction (Drmota and Kral 1997). Previous studies of T.
vaginalis karyotype by FISH showed that constriction on chromosome IV is secondary where
rRNA genes are localized (Torres-Machorro et al. 2009; Zubacova et al. 2011). Thus
subtelomeric constrictions on chromosomes I, III and V are most likely primary and
correspond to centromeres. The remaining chromosomes II, VI and also chromosome IV
without primary constrictions are classified as acrocentric with terminal position of
centromeres (Drmota and Kral 1997; Zubacova et al. 2011). We attempted to confirm the
position of centromeres on T. vaginalis chromosomes by cenH3 immunostaining.
Overexpressed cenH3 variant was detectable on condensed chromosomes only after slight
denaturation of chromosomes with 0,1% SDS. The fluorescence pattern was different in
comparison to the other variants but the morphology of metaphase chromosomes was not
preserved to identify the precise position of centromeres (Figure 3.).
Overexpression of neither core H3 nor its variants did not affect growth and viability
of Trichomonas cells. As cenH3 was found to be essential for cell survival (Cervantes et al.
2006; Lowell and Cross 2004), study of phenotype of T. vaginalis cenH3 gene knock-outs
remains attractive proposition. In T. vaginalis genome are annotated also several conserved
proteins of kinetochore complex, such as CENP – B or CENP – E (Table 1.), co - localization
experiments could support the role of histone TVAG_224460 as centromeric marker.
In conclusion, we found that T. vaginalis has H3.3 histone variant that is targeted to
transcriptionally active regions of chromatin. We showed that one of the two distinct H3
variants has function as epigenetic marker of centromeres. The function of the third
trichomonad H3 variant is unclear.
TVAG_129730
TVAG_224650
TVAG_100590
TVAG_399370
TVAG_341360
TVAG_414480
TVAG_211260
TVAG_085670
TVAG_499020
TVAG_137940
TVAG_497330
TVAG_445270
TVAG_452730
TVAG_315320
TVAG_470690
TVAG_270890
TVAG_014910
TVAG_270800
TVAG_239330
TVAG_270080
TVAG_087830
TVAG_185390
TVAG_224460
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKSTG---GKTPRKSLGAKAARKSTPTID-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MARTKQTARKTTG---GKTPRKSLGAKAARKAIPTVD-SQGAKKQHRFRPGTVALREIRKYQKSTDLLIR
MEEEPRPIHR---GKKRRIPLSSGAARPPDNSSDKSESQKKQRRKR--NSWLREIHFYQKTTNLLIR
MASTRIYADDWSFFDDPDNRSSRLEFSIPSQWSQLDVIKPAKKQLKKKKLPNPDQKPKKKHNHVLKEIRTYQNSVDLLIP
66
66
66
66
66
66
66
66
66
66
66
66
66
66
66
66
66
66
66
66
66
62
80
TVAG_129730
TVAG_224650
TVAG_100590
TVAG_399370
TVAG_341360
TVAG_414480
TVAG_211260
TVAG_085670
TVAG_499020
TVAG_137940
TVAG_497330
TVAG_445270
TVAG_452730
TVAG_315320
TVAG_470690
TVAG_270890
TVAG_014910
TVAG_270800
TVAG_239330
TVAG_270080
TVAG_087830
TVAG_185390
TVAG_224460
KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGERN-KLPFQRLVREIASGFR-GDLRFQSSAIAALQEAAEAYLVGLFEDTNLCAIHANRVTIMERDVQLAMRIRGERN-KLPFCRLVKEITQSVSIGEFRYTTGAMEALQEASEAFLIKLLEDGQVCAIHARRITLMNRDLQLAQRLRGDR--RLSFQRLVREIAHQNN-PTIKFQETAIQALQEASEAFLVGMMEDGNLCTIHAQRVTIMKKDMKLAERIRGDSITE
72
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74
Figure 1a. Sequence alignment of all histones H3 identified in T. vaginalis genome database TrichDB
(http://trichdb.org/trichdb/). Histones highlighted in red were used in experiments: TVAG_270080 as core H3;
TVAG_087830 as H3.3 variant; TVAG_185390 as H3 variant with unknown function and TVAG_224460 as
cenH3.
Histone fold domain
TVAG270080
TVAG224460
TVAG185390
SpoCnp1
HsCenp-A
GiCenH3
TbH3V
-MA-RTKQTARKSTGGKTPRKS-----LGAKAARKSTPTIDS-QGAKKQHRFRPGTVALREIRKYQKSTDL
-MASTRIYADDWSFFDDPDNRSSRLEFSIPSQWSQLDVIKPAKKQLKKKKL-PNPDQKPKKKHNH-VLKEIRTYQNSVDL
-MEEEPRPIHRGKKRRIPLSS-----GAARPPDNSSDK----SESQKKQRRKRNS-WLREIHFYQKTTNL
-MA-KKSLMAE--PGDPIPR------------PRK--------------KRYRPGTTALREIRKYQRSTDL
-MGPRRRSRKPEAPRRRSPSPT-----PTPGPSRRG-PSLG--ASSHQHSRRRQG--WLKEIRKLQKSTHL
-MSGGSRSQVARNTGHRRREISGRNMIPGVVVNARQSRSKLSSDPFSSVPRRPARVSHMEREIYHYQHNVDT
-MAQMKKITPR--PVRPKSVASR---P-IQAVARAPVKKVENTPPQKRHHRWRPGTVALREIRRLQSSTDF
63
77
59
41
60
71
64
Loop1
TVAG270080
TVAG224460
TVAG185390
SpoCnp1
HsCenp-A
GiCenH3
TbH3V
LIRKLPFQRLVREIASGFRG------DLRFQSSAIAALQEASEAYLVGLFEDTNLCAIHANRVTIMERDVQLAQRIRGER
LIPRLSFQRLVREIAHQNNP------TIKFQETAIQALQEASEAFLVGMMEDGNLCTIHAQRVTIMKKDMKLAERIRGDS
LIRKLPFCRLVKEITQSVSIG-----EFRYTTGAMEALQEASEAFLIKLLEDGQVCAIHARRITLMNRDLQLAQRLRGDR
LIQRLPFSRIVREISSEFVANFSTDVGLRWQSTALQCLQEAAEAFLVHLFEDTNLCAIHAKRVTIMQRDMQLARRIRGALIRKLPFSRLAREICVKFTRG----VDFNWQAQALLALQEAAEAFLVHLFEDAYLLTLHAGRVTLFPKDVQLARRIRGLE
LIQKLPFARLVQELVEQIAQRDGSKGPYRFQGMAMEALQSATEEYIVELFSTALLATYHANRVTLMSKDILLVLRIQ-QR
LIQRAPFRRFLREVVSNLKDS------YRMSAACVDAIQEATETYITSVFMDANLCTLHANRVTLFPKDIQLALKLRGER
TVAG270080
TVAG224460
TVAG185390
SpoCnp1
HsCenp-A
GiCenH3
TbH3V
N--------ITE------------------------EGLG-----NLNSLR---N---------
74
74
75
79
76
79
74
1
3
0
0
4
6
1
Figure 1b. Sequence alignment of T. vaginalis histone H3 variants (TVAG_270080 as core H3; TVAG_224460
as cenH3, TVAG_185390 as H3 variant with unknown function). Centromeric H3 variants cenH3s have
typically divergent N – termini that can not be aligned between species (yellow). In contrast to cenH3s of other
organisms, T. vaginalis cenH3 does not have substituted glutamine residue in the α-1 helix of the protein (pink)
and lacks the insertion in the loop 1 region (grey).
Abbreviations: Spo – Schizosaccharomyces pombe, Hs – Homo sapiens, Gi – Giardia intestinalis, Tb –
Trypanosoma brucei.
DAPI
á-HA
DAPI + á-HA
Canonical histone H3 TVAG_270080
Histone H3 variant TVAG_185390
H3.3 variant TVAG_087830
CenH3 variant TVAG_224460, G1 phase
CenH3 variant TVAG_224460, G2 phase
Figure 2. Localization of trichomonad core H3 and H3 variants during interphase.
DAPI
á-HA
DAPI+á-HA
Core histone H3 TVAG_270080
Histone H3 variant TVAG_185390
H3.3 variant TVAG_087830
CenH3 variant TVAG_224460
Figure 3. Localization of trichomonad core H3 and H3 variants during metaphase.
á-HA
mH3K4
merge
H3.3 variant TVAG_087830
Figure 4. Active regions of transcription visualised by monomethyl H3K4
immunostaining colocalize with T. vaginalis H3.3 variant TVAG_087830.
Annotation
Gene ID
Function in mitosis
cenp-A
TVAG_224460
DNA binding, centromere assembly
cenp-B
TVAG_067240
DNA binding, centromere assembly
inner centromere protein
TVAG_475090
chromosomal passenger protein, cohesion
inner centromere protein
TVAG_123230
chromosomal passenger protein, cohesion
inner centromere protein
TVAG_215610
chromosomal passenger protein, cohesion
cbf5, putative
TVAG_407130
chromosomal passenger protein
centromeric protein E
TVAG_237710
outer kinetochore, movements
centromeric protein E
TVAG_010490
outer kinetochore, movements
centromeric protein E
TVAG_494200
outer kinetochore, movements
centromeric protein E
TVAG_026780
outer kinetochore, movements
centromeric protein E
TVAG_410100
outer kinetochore, movements
centromeric protein E
TVAG_099890
outer kinetochore, movements
centromeric protein E
TVAG_244940
outer kinetochore, movements
centromeric protein E
TVAG_192230
outer kinetochore, movements
centromeric protein E
TVAG_473490
outer kinetochore, movements
centromeric protein E
TVAG_193720
outer kinetochore, movements
centromeric protein E
TVAG_030630
outer kinetochore, movements
centromeric protein E
TVAG_480170
outer kinetochore, movements
centromeric protein E
TVAG_458090
outer kinetochore, movements
centromeric protein E
TVAG_158440
outer kinetochore, movements
centromeric protein E
TVAG_007500
outer kinetochore, movements
centromeric protein E
TVAG_307050
outer kinetochore, movements
high mobility group protein
TVAG_402230
centromere assembly
high mobility group protein B2
TVAG_110110
centromere assembly
high mobility group protein B1
TVAG_485890
centromere assembly
high mobility group protein
TVAG_302620
centromere assembly
high mobility group protein
TVAG_453060
centromere assembly
high mobility group protein
TVAG_161660
centromere assembly
high mobility group protein
TVAG_325010
centromere assembly
high mobility group protein
TVAG_125540
centromere assembly
high mobility group protein
TVAG_194720
centromere assembly
high mobility group protein
TVAG_346400
centromere assembly
DNA topoisomerase II
TVAG_038880
chromosomal passenger protein, release
DNA topoisomerase II
TVAG_198840
chromosomal passenger protein, release
cytoplasmic dynein heavy chain
TVAG_365640
movements
cytoplasmic dynein heavy chain
TVAG_432890
movements
cohesin subunit rad21
TVAG_297540
cohesion
spindle assembly checkpoint component MAD1
TVAG_186470
spindle checkpoint proteins
spindle assembly checkpoint component MAD1
TVAG_077120
spindle checkpoint proteins
spindle assembly checkpoint component MAD1
TVAG_483050
spindle checkpoint proteins
spindle assembly checkpoint component MAD1
TVAG_054210
spindle checkpoint proteins
spindle assembly checkpoint component MAD1
TVAG_181280
spindle checkpoint proteins
spindle assembly checkpoint component MAD1
TVAG_023960
spindle checkpoint proteins
TVAG_340790 peri-centromeric
heterochromatin protein
Table 1. T. vaginalis centromere – associated proteins, kinetochore and spindle – checkpoint components
identifyied in genome database (http://trichdb.org/trichdb/, (Santaguida and Musacchio 2009).
Reference List
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of the National Academy of Sciences of the United States of America 99:16477-16484
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Peer Y, Miranda-Saavedra D, Barton GJ, Westrop GD, Muller S, Dessi D, Fiori PL, Ren QH,
Paulsen I, Zhang HB, Bastida-Corcuera FD, Simoes-Barbosa A, Brown MT, Hayes RD,
Mukherjee M, Okumura CY, Schneider R, Smith AJ, Vanacova S, Villalvazo M, Haas BJ,
Pertea M, Feldblyum TV, Utterback TR, Shu CL, Osoegawa K, de Jong PJ, Hrdy I,
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Dunne RL, Upcroft JA, Upcroft P, White O, Salzberg SL, Tang P, Chiu CH, Lee YS, Embley
TM, Coombs GH, Mottram JC, Tachezy J, Fraser-Liggett CM, Johnson PJ (2007) Draft
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Cervantes MD, Xi XH, Vermaak D, Yao MC, Malik HS (2006) The CNA1 histone of the ciliate
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Dawson SC, Sagolla MS, Cande WZ (2007) The cenH3 histone variant defines centromeres in Giardia
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Dernburg AF (2001) Here, there, and everywhere: Kinetochore function on holocentric chromosomes.
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5. Thesis summary
Karyotype of Trichomonas vaginalis
Trichomonads reproduce asexually by mitosis. Cultures of trichomonads can be
enriched in cells with condensed and well – structured chromosomes by arresting the cells in
mitotic metaphase using colchicine. In T. vaginalis, mitotic index (a ratio between cells in
mitosis and total cell number) after colchicine treatment was in all experiments high. About
80% cells displayed visible chromosomes. We studied chromosomes of T. vaginalis by
conventional cytogenetic technique using chromosome spreading after short time hypotonic
treatment followed by fluorescence staining. T. vaginalis has haploid number of six tiny
chromosomes (the longest chromosome I is 2, 4 µm long) that differ in size and morphology.
Each trichomonad chromosome consist of two chromatids that are arranged in parallels,
however, connection between sister chromatids typical for monocentric as well as holocentric
chromosomes of higher eukaryotes was either not visible or observed very rarely.
Constrictions were observed on chromosomes I, III, IV. and V. suggesting that the
chromosomes of T. vaginalis are monocentric with point centromeres. Chromosome number
in T. vaginalis karyotype is stable in different isolates (we tested strains T1, G3 and Cp1) and
does not vary after long term cultivation. We did not observed pairing of homologous
chromosomes typical for meiotic division. Our failure to observe meiosis in T. vaginalis,
however, does not exclude a possibility that this process may rarely occur.
Karyotypes of related trichomonads
In addition to Trichomonas vaginalis, we investigated and revised karyotypes in other
eight species including Trichomonas tenax (6 chromosomes), Tetratrichomonas gallinarum (5
chromosomes), Pentatrichomonas hominis (6 chromosomes), Tritrichomonas foetus (5
chromosomes), Tritrichomonas augusta (5 chromosomes), Monocercomonas colubrorum (4
chromosomes), Trichomitus batrachorum (6 chromosomes) and Hypotrichomonas acosta (5
chromosomes). Karyotypes of this species have not been studied previously or data about
their chromosomes are from the beginning of twenty century when the limits of light
microscopy could affect the results. All species can be studied by spreading technique using
colchicine treatment, however observed mitotic index was lower when compared to T.
vaginalis. All trichomonads have single chromosome set with constant number of
chromosomes. Sister chromatid organization without visible cohesion was the same as in T.
vaginalis.
42
Nuclear DNA content and genome sizes in related trichomonads
Genome sequence project of T. vaginalis revealed that this organism possesses the
largest genome among parasitic protists. It was hypothesized that the large size of the genome
is related with pathogenic potential of T. vaginalis. Thus, we analyzed nuclear DNA content
of other eight species from Trichomonadea group. T. vaginalis cells were used as standard
with known genome size of 160 Mb. Flow cytometry revealed relatively large genomes in all
tested species ranging from  90 Mb in human commensal Pentatrichomonas hominis to 177
Mb in tritrichomonads. Therefore, previously suggested relationship between large genome
size of trichomonads and parasitic life style, extraintestinal localization in host and
phagocytosis ability was not confirmed in our analysis. The only positive correlation was
found between genome size versus cell and nuclei volume in all tested trichomonads. Based
on inferred earliest emergence of hypermastigids among the Parabasala group, the giant protists
living in termite intestine, we suggested that unusually large genome of T. vaginalis may be
not the result of recent expansion but oppositely it resulted from regressive evolution of large
genome of trichomonad common ancestor.
Fluorescence in situ hybridization as a new tool in research of genome organization of
Trichomonas vaginalis
We tested classic cytogenetic techniques including C – banding and fluorescence
banding to find distribution of constitutive heterochromatin on T. vaginalis chromosomes but
no bands were observed using any technique. We have, however, adapted fluorescence in situ
hybridization (FISH) for T. vaginalis metaphase chromosomes that is sensitive enough to map
not only repetitive sequences but also single copy genes. We found markers for chromosome
I. and II. by mapping the localization of single copy genes encoding serine
palmitoyltransferase and tryptophanase. Previous attempts to detect nucleolar organizer
regions (NORs) on T. vaginalis by AgNO3 staining failed. FISH mapping of ribosomal RNA
genes revealed single NOR in secondary constriction of chromosome IV. FISH was
successfully employed in the study of genetic diversity of T. vaginalis strains of diverse origin
using panel of microsatellites as genetic markers. Analysis revealed the existence of genetic
variation among T. vaginalis strains and indicates participation of genetic recombination on
genetic diversity of trichomonads. Our FISH protocol provides a tool for physical mapping of
T. vaginalis genome as well as for development of additional genetic markers and study of
genome rearrangements.
43
Histone H3 variants in Trichomonas vaginalis and identification of centromeric marker
cenH3
In T. vaginalis genome database TrichDB there are 23 genes for histone H3. Twenty
of them are identical and according to the sequence alignments they are core histones H3. The
remaining three sequences are different. We investigated their protein sequences and nuclear
chromatin localization by transfection of trichomonads with hemagglutinin – tagged histone
constructs. We wanted to determine if they belong to known group of eukaryotic H3 variants
such as marker of transcriptionally active genes H3.3 and divergent centromere marker cenH3
responsible for proper chromosome segregation and to distinguish whether eitherer of these
histone H3 variants define trichomonad centromeres.
Generally, core histones represent one of the most conserved proteins with the
exception of histones from protists that show significant divergence. Overall diversity of core
H3 sequences of metazoans is typically less than 15%, however, differences between protist
and other organisms can be up to 44%. Comparison of T. vaginalis core H3 protein sequence
with homologs from other organisms revealed only 82% identity with sequences from
multicellular organisms (Dernburg, 2001; Marinets et al., 1996; Zubacova and Tachezy,
unpublished data).
Trichomonad putative H3.3 variant (TVAG_087830) is almost identical with protein
sequence of core H3. They differ only in six amino acid residues. This histone H3 variant co localized with active transcriptional regions in chromatin as defined by monomethyl H3K4
immunostaining that strongly suggest that this histone is H3.3 ortholog.
Two other divergent H3 variants in T. vaginalis TVAG_185390 and TVAG_224460
have different N – terminal parts of the protein and can be aligned with core H3 only in their
histone fold domain where sharing 60% identity. H3 variant TVAG_185390 is unique for T.
vaginalis. Although it shows some sequence features of cenH3 including amino acid insertion
in loop 1 region and absence of conserved glutamine residue in alpha – 1 helix in protein
structure, its deposition in chromatin was congruent with core H3 histone. The role of this
variant
in
trichomonad
chromatin
44
is
not
known.
The second variant TVAG_224460 named TvcenH3 is most likely cenH3 ortholog
according to specific immunostaining pattern. It localized to six and twelve discrete foci,
corresponding to chromosome number of T. vaginalis in G1 and G2 phase, respectively. The
sequence alignment of trichomonad core histones H3 and H3 variants revealed short N –
terminal extension of two amino acids in TvcenH3 that is a feature of all studied cenH3s.
Protein sequence of TvcenH3, however, seems to be unique as it lacks insertion in loop 1 and
has conserved glutamine in alpha – 1 helix. As T. vaginalis cenH3 is encoded by single copy
gene, it would be interesting to study phenotype of cenH3 knock – outs to test the effect on
cell viability because cenH3 is considered to be essential protein for cell division.
Together with cenH3 we have identified also several genes coding conserved
kinetochoral proteins in T. vaginalis genome such as CENP-B, CENP-E or INCENP
suggesting that segregation of T. vaginalis chromosomes is mediated by conventional
attachment of spindle microtubules to kinetochores. Localization of trichomonad putative
kinetochoral components as well as visualization of chromosomes capture by extranuclear
spindle microtubules in T. vaginalis closed mitosis remains a subject of future studies.
45