AtHaspin phosphorylates histone H3 at threonine 3 during mitosis and contributes to embryonic patterning in Arabidopsis

AtHaspin phosphorylates histone H3 at threonine 3 duringmitosis and contributes to embryonic patterning inArabidopsis

Raheleh Karimi Ashtiyani, Ali Mohammad Banaei Moghaddam, Veit Schubert, Twan Rutten, Jorg Fuchs, Dmitri Demidov,

Frank R. Blattner and Andreas Houben*

Leibniz Institute of Plant Genetics and Crop Plant Research, Corrensstr. 3, 06466 Gatersleben, Germany

Received 26 May 2011; accepted 25 June 2011; published online 30 August 2011.

*For correspondence (fax +49 394 82 5137; e-mail [email protected]).

SUMMARY

Post-translational histone modifications regulate many aspects of chromosome activity. Threonine 3 of

histone H3 is highly conserved, but the significance of its phosphorylation is unclear, and the identity of the

corresponding kinase in plants is unknown. Therefore, we characterized the candidate kinase in Arabidopsis

thaliana, called AtHaspin. Recombinant AtHaspin in vitro phosphorylates histone H3 at threonine 3. Reduction

of H3 threonine 3 phosphorylation level and reduced chromatin condensation in interphase nuclei by AtHaspin

RNAi supports the proposition that this kinase is involved in histone H3 phosphorylation in vivo in mitotic

cells. In addition, we provide a developmental function for a Haspin kinase. At the whole plant level, altered

expression of the kinase induced pleiotropic phenotypes with defects in floral organs and vascular tissue. It

reduced fertility and modified adventitious shoot apical meristems that then gave rise to plants with multi-

rosettes and multi-shoots. Haspin mutant embryos frequently showed alteration in division plane orientation

that could be traced back to the earliest divisions of embryo development, thus Haspin contributes to

embryonic patterning.

Keywords: embryo development, haspin, histone H3 phosphorylation, kinase, mitosis.

INTRODUCTION

The cell cycle-dependent transition from decondensed

interphase chromatin to condensed metaphase chromatin,

and vice versa, is accompanied by the phosphorylation of

histones. Histone H3 is specifically phosphorylated during

both mitosis and meiosis, but also plays a role in gene

activation and stress regulation (reviewed in Hans and

Dimitrov, 2001; Houben et al., 2007). It is noteworthy that

although cell cycle-dependent phosphorylation at different

histone H3 amino acid residues is conserved between plants

and non-plants, the functional significance of post-transla-

tional histone modification apparently changed during

evolution (Loidl, 2004; Fuchs et al., 2006; Houben et al.,

2007).

Phosphorylation at histone H3 threonine 3 (H3T3) is

accompanied by the condensation of chromosomes during

mitosis in vertebrates (Polioudaki et al., 2004; Dai et al.,

2005) suggesting a functional role for this post-translational

modification in chromosome condensation and segrega-

tion. The enzyme identified for the cell cycle-dependent

phosphorylation at histone H3T3 of non-plant organisms is

the serine/threonine kinase Haspin (reviewed in Higgins,

2010).

Haspin (haploid cell-specific protein kinase) was first

discovered in male germ cells of mice (Tanaka et al., 1999).

This kinase is most strongly expressed in testis, but is also

appears ubiquitous in proliferating somatic cells, including

human tumor lines (Dai et al., 2009; Markaki et al., 2009).

Microscopical detection of myc- or GFP-tagged Haspin

revealed a cell cycle-dependent association with condensed

chromosomes and centrosomes throughout mitosis

(Tanaka et al., 1999; Dai and Higgins, 2005; Dai et al., 2005).

Crystal structure analysis of the kinase domain of human

Haspin confirms that it forms a bilobed structure similar to

that of eukaryotic protein kinases (ePKs), but with significant

structural changes and a number of Haspin-specific inserts

(Eswaran et al., 2009; Villa et al., 2009). Haspin immuno-

precipitated from cells co-purifies with and phosphorylates

histone H3, and recombinant Haspin specifically phospho-

rylates purified and nucleosomal histone H3 in vitro at

threonine 3 (Dai et al., 2005; Patnaik et al., 2008; Eswaran

ª 2011 The Authors 443The Plant Journal ª 2011 Blackwell Publishing Ltd

The Plant Journal (2011) 38, 443–454 doi: 10.1111/j.1365-313X.2011.04699.x

et al., 2009; Villa et al., 2009). The ectopic expression of

mammalian Haspin led to an increased level of H3T3

phosphorylation, delayed mitosis and reduced proliferation

(Dai et al., 2005). In contrast, depletion of Haspin by RNA

interference (RNAi) resulted in a reduced level of mitotic

H3T3 phosphorylation and prevented normal chromosome

alignment at metaphase (Dai et al., 2005).

In addition, human Haspin interacts with cohesin, a

protein complex required for sister chromatid cohesion,

and regulates chromosome segregation during mitosis

together with other proteins which are required for the

maintenance or removal of chromosomal cohesion, such as

Sgo1, Bub1, CENP-F, Aurora B, and Plk1 (Dai et al., 2006,

2009; Rosasco-Nitcher et al., 2008; Yamagishi et al., 2010).

Recent studies have revealed that phosphorylation of

histone H3T3 by Haspin is necessary for recruitment of the

chromosome passenger complex components Aurora B and

Survivin (Kelly et al., 2010; Wang et al., 2010).

Haspin homologues have been identified in silico for a

number of eukaryotic lineages, including yeast and plants

(Higgins, 2001). The existence of at least one Haspin

homologue in all nearly completely sequenced eukaryotic

genomes suggests an important function, and an apparent

early origin for this protein (Higgins, 2003; Nespoli et al.,

2006).

A cell cycle-dependent post-translational modification of

histone H3T3 has also been demonstrated for higher plants

(Houben et al., 2007). Interestingly, the results confirm prior

suggestions that the distribution of phosphorylation on

histone H3 of mitotic chromosomes is almost reversed in

plants compared with mammals. While phosphorylation of

H3 at serine 10 (H3S10) is restricted to pericentromeric

regions in plants, H3T3ph originates at pericentromeres in

prophase and is evenly distributed along chromosome arms

by prometaphase (Houben et al., 2007; Caperta et al., 2008).

The reasons for these differences are unknown, but it

appears that there has been considerable divergence in the

roles of histone modification between plants and other

organisms (Loidl, 2004; Houben et al., 2007). Phosphoryla-

tion of histones by the recombinant budding yeast Haspin

homologues Alk1 and Alk2 was not observed in vitro

(Nespoli et al., 2006).

Although plants possess Haspin homologues, it remains

to be determined whether its function is conserved in

evolution and also whether it phosphorylates histone H3 at

T3. In addition, as most studies on Haspin were conducted

only on cell lines, the contribution of Haspin to the devel-

opment of a complex organism remains to be determined.

In this report we characterized in A. thaliana the corre-

sponding Haspin-like kinase (called AtHaspin) responsible

for the phosphorylation of histone H3 at threonine 3. Partial

inactivation of AtHaspin by RNA interference reveals that it is

involved in histone H3T3 phosphorylation in mitotic cells

and condensation of interphase chromosomes. At the whole

plant level, altered expression of the kinase induced pleio-

tropic phenotypes, which included reduced fertility, adven-

titious shoot apical meristem formation and abnormal

embryo development. Our work reveals a new kinase

involved in composing the histone code in plants and adds

AtHaspin to the select group of genes that contributes to

pattern formation in the A. thaliana embryo, as indicated by

the mutant phenotype.

RESULTS

Plants encode a conserved Haspin-like protein

By BLASTX analysis of the non-redundant nucleotide

sequence database of the A. thaliana genome using the

amino acid sequence of the catalytic domain of the human

Haspin we could identify a Haspin-like serine/threonine

kinase (called AtHaspin), which is encoded by the locus

At1g09450. AtHaspin contains 15 exons separated by 14

introns in approximately 4 kb of coding regions that encode

a protein of 599 amino acids with a predicted molecular

weight of approximately 68 kDa (Figure S1).

Alignment of the Haspin-kinase domains of plants, human

and mouse allowed the identification of the eukaryotic

protein kinases (ePKs) typical subdomains I to XI (Higgins,

2001). Many of the residues that are essentially invariant in

other ePKs, and known to be critical in formation of the

Mg2+-ATP-binding and catalytic sites (Higgins, 2001) are

conserved in the majority of Haspin proteins (Figure S2).

Specifically, the G-x-G-x-x-G-x-(V/A) motif in region I, the

lysine (K) in region II, glutamate (E) in region III, aspartic acid

(D) and asparagine (N) in region VIb, aspartic acid (D) in

region VII, and aspartic acid (D) or glutamate (E) in region IX

are conserved. Plant-specific sequence motives of the kinase

domain are indicated in Figure S2.

Haspin-like proteins have been identified in all eukaryotic

genomes so far sequenced (Higgins, 2001, 2003). To inves-

tigate the evolutionary relationship of the Haspin family in

plants, the conserved catalytic domain was used to build a

phylogenetic tree (Figure S3). We searched the EMBL plant

EST database, and queried with the predicted protein

sequences of AtHaspin. Cladistic (MP) analysis of the amino

acid alignment of Haspin kinases resulted in 30 most

parsimonious trees (length 7678 steps, consistency index =

0.520, retention index = 0.547). The strict consensus tree of

this analysis is completely compatible with the result of the

phylogenetic analysis (NJ). Therefore, only the NJ tree is

given (Figure S3).

In higher plants mainly one type of Haspin kinases seems

to be present, as all sequences form a monophyletic group

that is sister to the moss Physcomitrella patens, and both

groups together are a sister group of the algae genus

Ostreococcus. Sequence relationships within angiosperms

reflect the phylogeny of this group with a split between

sequences derived from monocot and eudicot taxa. In

444 Raheleh Karimi Ashtiyani et al.

ª 2011 The AuthorsThe Plant Journal ª 2011 Blackwell Publishing Ltd, The Plant Journal, (2011), 38, 443–454

contrast, in Brugia malayi, Saccharomyces cerevisiae and

Caenorhabditis elegans paralogous Haspin kinases occur.

The sequences derived from Placozoa (Trichoplax), insects

and vertebrates are monophyletic and occur together with

plants and the Choanoflagellate Nonosiga brevicollis in an

unresolved polytomy, while one group of sequences derived

from the nematodes Caenorhabditis and Brugia form the

sister group to all these organisms. The other sequences

derived from C. elegans are quite diverse, and separated

from the clades already described by all sequences derived

from fungi.

AtHaspin phosphorylates histone H3 at threonine 3 in vitro

To determine whether the phosphorylation activity of Haspin

towards threonine 3 of histone H3 is conserved during plant

evolution, we tested the ability of recombinant AtHaspin to

phosphorylate histone H3T3. The activity of the recombinant

AtHaspin was analyzed using an in vitro kinase assay with

two different substrates, namely total histone extracts,

including H3 isolated from calf thymus, and myelin basic

protein (MBP) (Figure 1a). The in vitro kinase assay demon-

strated that, like the human Haspin protein (Dai et al., 2005),

the recombinant AtHaspin shows protein kinase activity

towards total histone H3 (Figure 1a, lane 4). In addition, a

weaker kinase activity towards other histones was also

observed. The phosphorylation signal was absent when

AtHaspin was omitted from the reaction mixtures (Figure 1a,

lane 3), or in the reaction contained AtHaspin only (Figure 1a,

lane 2). Unlike the findings in mammals (Tanaka et al., 1999;

Dai et al., 2005), recombinant Haspin of A. thaliana has a

weak autophosphorylation capability whether or not histone

H3 or other substrates are present (Figure 1a), while no

phosphorylation was observed when we used vector alone

(Figure 1c). In parallel, we prepared a control Haspin protein

(AtHaspin-mut) that contained a mutation of a single con-

served lysine residue (K310A) in the catalytic domain of the

enzyme (Figure 1c). The in vitro kinase assay using AtHa-

spin-mut demonstrated that the lysine 310 residue is

essential for enzyme activity, as reported for other protein

kinases (Dai et al., 2005). Next, we determined the amino

acid residue of histone H3 that is phosphorylated by AtHa-

spin. Non-radioactive in vitro kinase assays (Figure 1d) were

performed with protein extracts from recombinant AtHaspin

using total histone H3 as a substrate. Subsequent immuno-

blot analyses demonstrated a specific phosphorylation at T3

of histone H3. No significant phosphorylation was detected

at the histone H3 sites serine 28, serine 10 or threonine 11.

Hence, the phosphorylation activity of Haspin towards thre-

onine 3 of histone H3 is conserved during evolution.

AtHaspin is highly expressed in tissues with a high level of

cell proliferation and differentiation

To analyze contribution of Haspin to the development of a

complex organism the tissue type-specific expression pat-

tern was analyzed first. Expression patterns of AtHaspin

determined by semi-quantitative reverse transcription

polymerase chain reaction (RT-PCR; Figure 2a) and in

silico analysis (Genevestigator, https://www.genevestigator.

com/gv). Figure 2b) showed the highest abundance of

transcripts in proliferating and differentiating organs, such

as flower buds and flowers, as well as stem-bearing shoot

meristems. Low levels of transcription were found in other

organ types. Because AtHaspin mRNA was abundant in

extracts from tissue rich in dividing cells, we plotted the RNA

profiles of AtHaspin in synchronized A. thaliana tissue

culture cells using publicly available microarray datasets

(Menges et al., 2002; Figure 2c). In contrast to the cell cycle-

dependent transcription behavior of cyclin B, AtHaspin

transcription does not change significantly during the

mitotic cell-cycle.

Figure 1. Phosphorylation of histone H3T3 by recombinant AtHaspin.

(a) In vitro kinase reactions of recombinant 6xHis-AtHaspin with or without

the addition of total histone mix and myelin basic protein (MBP) as substrates.

6XHis-AtHaspin shows kinase activity toward total histone mix (a, lane 4) and

MBP (a, lane 1). The phosphorylation signal was absent when AtHaspin (a,

lane 3) or substrates (a, lane 2) was omitted from the reaction mixtures.

(b) The positions of histones and 6XHis-AtHaspin in the gel were determined

by Coomassie blue staining.

(c) In vitro kinase reaction of recombinant 6XHis-AtHaspin, 6XHis-AtHaspin-

mut and vector alone in the presence of total histone mix. No phosphorylation

was detected when we used recombinant protein for 6XHis-AtHaspin-mut and

vector alone.

(d) Non-radioactive in vitro kinase assay performed with protein extracts from

recombinant AtHaspin using total histone mix as substrate and without

substrate as negative control. Western blot experiment using different

antibodies against histone H3 (anti-H3, as control), phosphorylated histone

H3 at T3, Ser10, T11 and Ser28 revealed specific phosphorylation at T3 of

histone H3. The arrow indicates the band corresponding to the position of

histone H3.

Arabidopsis Haspin-like kinase 445


In order to confirm the expression pattern of AtHaspin we

constructed promoter–GUS fusions using 1500-bp and 705-

bp long genomic fragments upstream of the start codon of

the AtHaspin gene to the GUS reporter gene. The application

of two different sized fragments allowed us to discriminate

possible effects of the 5¢ neighbor gene, as AtHaspin has a

1600-bp long common intergenic region with the gene

At1g09440, which is in reverse orientation with the AtHa-

spin, and predicted to be the promoter region of both genes

(Shahmuradov et al., 2005).

No differences were observed between plants transformed

with the 1500-bp or the 705-bp long promoterAtHaspin::GUS

constructs regarding the localization and intensity of GUS

signals. The strongest GUS expression was observed in the

shoot apex and in the vascular tissue of young leaves, but not

in the vascular tissues of the roots (Figure 3a,b,d). In inflo-

rescences expression was detected in the vascular system of

the pedicles (Figure 3c), developing sepals (Figure 3f) and

stamens (Figure 3f,j), and in the basal part of pistils (Fig-

ure 3e,f,g). AtHaspin is also expressed in the funiculus of

developing ovules (Figure 3h) and in cotyledon primordia of

mature embryos (Figure 3i). Consistent with the expression

analysis (Figure 2a,b) mature plants displayed a lower

expression of promoterAtHaspin::GUS in the vascular tissue

of all major aerial organs including stems, rosettes, flowers,

siliques. No promoterAtHaspin::GUS activity was found in

roots, which could be explained by the absence of putative

root-specific regulatory elements, which were not included in

promoterAtHaspin::GUS constructs. Arabidopsis plants trans-

formed with the empty pMDC162 vector did not reveal any

detectable GUS signal (Figure 3k).

Complete inactivation of AtHaspin is lethal for plants

and affects the cell division of the zygote

Four different A. thaliana lines in the Col-0 background with

T-DNA insertions in the promoter region or introns of the

AtHaspin gene were identified in the SALK (Alonso et al.,

2003) and GABI (Rosso et al., 2003) collections (Figure S1).

These lines were has1-1 for SALK_019798, has1-2 for GABI

435H08, has1-3 for GABI 082D07, and has1-4 for GABI

858F01. PCR-based genotyping of segregating pools of

plants revealed that we could obtain plants homozygous for

the T-DNA insertions in has1-1, has1-2 and has1-4.

Semi-quantitative RT-PCR of homozygous T-DNA plants of

has1-1, has1-2 and has1-4 did not reveal any AtHaspin

expression differences between homozygous T-DNA

mutants, and wild-type plants. The reason, why the tran-

scription behavior of AtHaspin was unaffected may be found

in T-DNA insertion positions, which were located either in

the 5¢ region upstream of the start codon (has1-2) or in intron

regions (has1-1 and has1-4).

There were also no obvious phenotypic differences

between wild-type and T-DNA plants. PCR analysis and

DNA sequencing of the PCR products confirmed that has1-3

contains a T-DNA insertion in the 5¢ untranslated region

(UTR) close to the start codon ()27 relative to the ATG start

codon). It was also notable that no homozygous T-DNA plant

for the AtHaspin gene was found in the progeny of hetero-

zygous has1-3 plants (Table S1). The observed segregation

ratio of the progeny of 94 self-pollinated heterozygous has1-3

plants (Table S1) showed 33% wild-type, 67% heterozygous

and no homozygous plants, (coupled with the 1:2:0 ratio)

indicating embryo lethality for complete inactivation of

AtHaspin. About 25% of immature seeds from heterozygous

has1-3 mutant plants were distinguishable by a white or

brown seed color (Figure 4a). Embryos of abnormal seeds

Figure 2. Comparative transcription analysis of AtHaspin.

(a) Semi-quantitative reverse transcription polymerase chain reaction (RT-

PCR) analysis of AtHaspin expression in different A. thaliana organs. Elonga-

tion factor 1B (EF-1B) was used as a control.

(b) AtHaspin expression behavior deduced from microarray data (http://

www.GeneVestigator.ethz.ch).

(c) mRNA profiles of AtHaspin during the mitotic cell cycle deduced from

publicly available microarray data of synchronized A. thaliana suspension

cells (Menges et al., 2003). While cyclin Cycb1 is activated at the onset of

mitosis, 8 h after release from the aphidicolin block, AtHaspin transcription

does not show any changes during mitotic cell cycle.



were arrested at the early globular stage of development

(Figure 4c). In contrast, green seeds of the same silique had

already reached the mature cotyledon stage (Figure 4a).

Whole-mount analysis of abnormal has1-3 seeds revealed

an aberrant pattern of development for suspensor cells and

embryos (Figure 4c). Unlike wild-type zygotes [reviewed in

Jenik et al. (2007)], which divide into large basal and small

apical cells (Figure 4b1) in has1-3 mutants, the plane of the

first cell division is skewed (Figure 4c1, arrowed). In wild-

type, the apical cell divides twice longitudinally and once

transversely to form the eight cells embryo (Figure 4b) and

the basal cell continued to divide vertically to produce

hypophysis and suspensor (reviewed in Jenik et al., 2007).

Has1-3 embryos show extra cell divisions in the hypophysis

(Figure 4c2,c3,c5) and suspensor cells (Figure 4c5). In normal

embryos, the hypophysis divides asymmetrically to produce

a lens-shaped cell at the globular stage, which is the

progenitor of the root tip (Figure 4b3), while in has1-3

embryos the hypophysis forms a cluster of cells (Figure

4c2,c5). As consequence of aberrant cell divisions embryos

at globular stage lack the characteristic cell arrangement and

shape (Figure 4c5,c6). This observation suggests that the

AtHaspin gene is required before the asymmetric division of

the zygote occurs. Hence, mutation of AtHaspin results in

abnormal embryo patterning, and it also indicates that

AtHaspin is an essential gene in A. thaliana.

Altered expression of AtHaspin results in pleiotropic

plant growth phenotypes

As the analysis of the T-DNA Arabidopsis insertion lines did

not result in the identification of viable adult knockout

plants, we reduced the level of AtHaspin transcript by RNA

interference (RNAi). The AtHaspin-RNAi construct employed

contains a 400 bp long AtHaspin-specific gene fragment

(Hilson et al., 2004). Semi-quantitative RT-PCR was used to

identify RNAi plants with reduced AtHaspin transcript level

(Figure 5a). These plants showed pleiotropic developmental

defects that included adventitious shoot apical meristem

formation (Figure 5e, arrowed) giving rise to several rosettes

(Figure 5d) and secondary shoot formation (Figure 5c).

(a)

(d)

(f) (g)

(j) (k)

(h) (i)

(b) (c)

(e)

Figure 3. PromoterAtHaspin::GUS localization in

different tissues of A. thaliana. In young seed-

lings (a, b, d) GUS was mainly localized in the

vascular bundle of the cotyledons, young leaves

and the shoot apical meristem and leaf primordia

(arrows). In inflorescences promoterAtHaspin::GUS

signals were localized to the pedicles (c), vascular

system of developing sepals (f) and stamens (f, j).

In the pistil, expression was detected in the

vascular system and strongly in the bases of the

style (g, e) and in the funiculus of the fertilized

ovules (h). During the embryogenesis pro-

moterAtHaspin::GUS is localized to the cotyledon

primordia (i). The empty GUS vector as negative

did not show any GUS activity (k). Scale bars

represent 500 lm.

Figure 4. AtHaspin is essential for correct embryo development.

(a) Immature siliques containing seeds in wild-type (WT) and heterozygous

has1-3 plants. About 25% of immature seeds from heterozygous has1-3

mutant plants are marked by white or brown seed color.

(b, c) Whole-mount analysis of embryo in wild- type plants (b1–b6) and

heterozygous has1-3 (c1–c6). Abnormal cell divisions in two cell embryo (c1)

and embryo proper (c4–c6) are indicated by arrows. In hypophysis (c2, c3, c5)

and suspensor (c5) extra cell divisions are marked by arrowheads. Scale bar

equals 25 lm.



Furthermore inflorescences and flower morphology were

also abnormal and fertility was often reduced (Figure 5f,g).

Out of 110 RNAi plants, 34% showed simultaneous forma-

tion of additional rosettes and shoots.

Similar abnormal growth phenotypes were observed in

plants expressing AtHaspin under the control of the constit-

utive cauliflower mosaic virus (CaMV) 35S promoter. Plants

with enhanced AtHaspin transcription activity (Figure 5b)

also showed multiple vegetative shoot apexes (Figure 5h,i).

Most of the vegetative meristems produced an inflorescence

stem, and consequently a multi-shoots phenotype. Thirty-

one out of 90 T1 AtHaspin overexpression plants displayed a

multi-rosette and multi-shoots phenotype.

The transcription activity of AtHaspin in the vascular

tissues led us to compare the leaf venation of wild-type and

AtHaspin-RNAi cotyledons. The vein patterning in wild-type

cotyledons is strongly constant with one primary vein that is

continuous with the hypocotyl vasculature, and typically

four secondary veins that branch from the primary vein

(Deyholos et al., 2000) (Figure 6a). In AtHaspin-RNAi seed-

lings, we frequently observed an aberrant and undeveloped

venation (Figure 6b–f). Apparently, down-regulation of

AtHaspin interferes with the development of the vascular

system in cotyledons.

The observed axillary shoot meristems formation, defects

in vascular tissue patterning, axillary buds and lateral organ

outgrowth, superficially resembles with phenotypes caused

by altered expression of genes acting on auxin transduction

pathway such as PIN1 (Okada et al., 1991), PIN6 (Petrasek

et al., 2006) or PID (Bennett et al., 1995). To test a potential

interplay, we investigated the effect of AtHaspin alteration on

the transcription level of some genes that are involved in the

transport of auxin (PIN1, PIN6 and PID) and recycling of auxin

transport component [GNOM (Mayer et al., 1993; Geldner

et al., 2003), GNOM-like (Nakano et al., 2009)] by quantitative

(q)RT-PCR. We found that in AtHaspin-RNAi plants the

expression of PIN1 and PID were decreased significantly

(Figure S4a). An almost similar tendency was found for plants

overexpressing AtHaspin for PIN6 and PID (Figure S4b).

However the transcription level of GNOM and GNOME like

did not significantly alter in the AtHaspin mutants. Thus,

activity of AtHaspin influences directly or indirectly the

transcription of genes involved in auxin transport pathway.

Down-regulation of AtHaspin affects chromosomal

histone H3T3 phosphorylation

To understand whether an altered activity of AtHaspin

influences the dynamics of the cell cycle-dependent

(a)

(c)

(h) (i) (j)

(d) (f)

(e)

(g)

(b) Figure 5. Analysis of plants overexpressing or

downregulating AtHapsin.

(a) Reverse transcription polymerase chain reac-

tion (RT-PCR)-based gene expression analysis of

AtHaspin RNAi; and (b) overexpressed plants

revealed altered expression of AtHaspin gene.

Plants with a reduced expression activity of

AtHaspin are marked with asterisks [in (a)].

Elongation factor 1B-specific primers were used

as control.

(c) AtHaspin-RNAi plants showing multi-shoots

phenotype, (g) semi-fertile and undeveloped

siliques, and (f) abnormal flowers (indicated with

arrows) when compared with (j) wild-type (WT)

plants.

(d) Homozygous AtHaspin-RNAi lines illustrate

multi-rosettes and multi-shoots formation.

Arrows indicate the position of adventitious

shoot meristems and lateral rosettes.

(h, i) Similar multi-rosettes phenotype was

observed in AtHaspin overexpressed plants.

(e) Adventitious vegetative shoot apical meris-

tems are indicated with arrows.

(i) Adventitious meristems are formed on the

primary inflorescence of AtHaspin overexpres-

sion mutants. Scale bar (g) represent 2 cm.



phosphorylation of histone H3T3, we performed immuno-

staining experiments on mitotic and meiotic cells obtained

from flower buds of AtHaspin down-regulation (AtHaspin

RNAi) using an antibody that recognised histone H3 phos-

phorylated at T3 (H3T3ph). In dividing cells of wild-type

plants H3T3 phosphorylation occurred along the entire

length of chromosomes and colocalized with the position of

condensed meiotic (metaphase I) and mitotic chromosomes

(Figure 7) as reported previously (Caperta et al., 2008). His-

tone H3T3 phosphorylation level was clearly reduced in

homozygous AtHaspin-RNAi plants (Figure 7). Here the

H3T3 phosphorylation signals were more restricted to the

pericentromeric regions of condensed chromosomes in

both mitotic and meiotic cells (Figure 7). Thus, indirect evi-

dence exists, that AtHaspin is involved in the cell cycle-

dependent phosphorylation of histone H3T3 in plants.

However, overexpression of AtHaspin did not alter the level

of histone H3T3 phosphorylation (Figure S5). Also, no

immunofluorescence signals were found in interphase cells

of overexpressing lines.

AtHaspin does not influence sister chromatid cohesion but

weakly chromatin condensation in interphase nuclei

The cell cycle-dependent phosphorylation of histone H3T3

correlates with the process of chromosome condensation

and segregation in plants (Houben et al., 2007; Caperta

et al., 2008). Therefore, we analyzed by fluorescent in-situ

hybridization (FISH) whether overexpression or down-

regulation of AtHaspin might influence condensation and

sister chromatid cohesion of chromosomes. As the

determination of the chromosome condensation degree in

cycling Arabidopsis cells is limited due to the size of

mitotic chromosomes we compared the degree of chro-

mosome condensation at interphase between plants with

a different transcription activity of AtHaspin. To assay the

influence of AtHaspin on the cohesion of sister chromat-

ids the frequency of interphase sister chromatid separa-

tion was assayed. In higher plants sister chromatids in

differentiated interphase nuclei are frequently separated

along euchromatic regions (Schubert et al., 2006, 2007,

2008).

In each case 4C leaf nuclei of five different 35S

overexpressing AtHaspin as well as RNAi transformants

were analyzed compared with wild-type using a centro-

mere-specific sequence (pAL) (Martinez-Zapater et al.,

1986), a contig of BACs to label chromosome 1 top

arm (CT1top; Schubert et al., 2008) and single BACs

(approximately 100 kb of T2P11, F11P17 and T1F9) to

identify chromosome 1 mid-arm positions (Figure 8a–d).

Figure 7. Immunodetection of histone H3 phosphorylated at threonine 3 in

meiotic and mitotic cells. The level of histone H3T3 phosphorylation is

reduced in AtHaspin-RNAi meiotic and mitotic cells compared to wild-type

(WT) DNA was counterstained with 4¢,6-diamidino-2-phenylindole (DAPI). The

merged picture shows DAPI in blue and phosphorylated H3T3 in red.

(a) (b)

(d)

(f)

(e)

(c)

Figure 6. Vascular defects in AtHaspin-RNAi plants. Vascular patterning in

cotyledons of (a) wild-type (WT) and (b–e) different RNAi plants.

(f) Diagram illustrates percentage of vascular pattern complexity in WT

(hatched bars) and RNAi plants (black bars) cotyledon. Scale bar indicates

0.5 mm.



Compared with wild-type plants positional sister chromatid

separation was decreased significantly in RNAi plants at

the top arm BAC position T2P11, but not at the bottom arm

position F11P17/T1F9. No significant difference between

wild-type (WT) and 35S plants was detected. At centro-

meres 0.5% (n = 1074) of RNAi and 0.7% (n = 1130) of

35S::AtHaspin nuclei, respectively, showed centromere

signal numbers >10. However, this amount did not differ

significantly from WT at P < 0.05. In short, our data do not

indicate a definite influence of AtHaspin on sister chroma-

tid cohesion in interphase nuclei. In contrast, the signifi-

cant chromatin condensation decrease of CT1top in RNAi

plants suggested that AtHaspin is involved in realizing

chromosome condensation along chromosome arms. At

centromeres no such significant effect on the condensation

behavior has been proven.

DISCUSSION

Histone H3T3 phosphorylation activity of Haspin is con-

served in plants

Recombinant AtHaspin in vitro phosphorylates histone H3 at

threonine 3. The same amino acid residue specificity has

also been demonstrated for mammalian Haspin (Dai et al.,

2005) but not for yeast Alk1 and Alk2 (Nespoli et al., 2006). In

contrast with Alk1 and Alk2 of yeast (Nespoli et al., 2006) and

Haspin of mammals (Tanaka et al., 1999; Dai et al., 2005)

AtHaspin of A. thaliana reveals only weak autophosphory-

lation activity.

Our promoterAtHaspin::GUS localization and transcription

data revealed the highest transcription activity of AtHaspin

in tissue types characterized by a high cell proliferation and

differentiation rate, which is consistent with the proposed

(a) (c)

(b) (d)

Figure 8. Sister chromatid cohesion and chromatin condensation in differentiated 4C interphase nuclei of 35S overexpressing and down-regulated (RNAi) AtHaspin

transformants, respectively, compared with A. thaliana wild-type (WT).

(a) Chromosomal location of fluorescence in-situ hybridization (FISH) probes detecting top arm of chromosome 1 (CT1top), centromeric 178-bp repeats (pAL) and

approximately 100 kb mid-arm segments (BACs T2P11, F11P17, T1F9). Significantly increased chromatin dispersion of CT1top was observed in RNAi plants. No

increased sister chromatid separation was present at both BAC loci of 35S and RNAi transformants. Left tables show association and dispersion frequency of top arm

of chromosome 1 (CT1top) and of the centromeric region (pAL). Right tables indicate frequency of sister chromatid separation at arm position T2P11, F11P17 and

T1F9. Number of 4C nuclei or homologous chromosomes analyzed are shown in parentheses.

(b) Nearly identical frequencies of centromere-specific FISH signals in AtHaspin and wild-type (WT) nuclei. Number of nuclei analyzed are shown in parentheses.

(c) Examples of RNAi nuclei labeled with BAC T1F9 (in red), the centromere-specific sequence pAL (in green) and counterstained with DAPI. T1F9 shows positional

separation at both chromosome 1 homologues. Seven centromeric signals indicate sister centromere cohesion and association (top) whereas the 16 signals of the

bottom nucleus refer to sister centromere separation.

(d) Positional sister chromatid cohesion and association of T2P11 (in green) visible within both associated compact CT1top (in red) homologues of a 35S AtHasin

nucleus (top). Bottom: CT1top chromatin decondensation and positional mid-arm sister chromatid separation at both homologues of an RNAi nucleus (left). Two

compact separated CT1 top arms showing cohered and separated T2P11 BAC positions (right).



cell cycle-dependent phosphorylation of histone H3T3 by

AtHaspin. But, unlike mitotic kinases, such as AtAuroras,

that are cell cycle-dependent transcriptionally active (Dem-

idov et al., 2005; Kawabe et al., 2005), plant (our data) and

also human Haspin (Dai et al., 2005) is expressed at near

constant levels throughout the cell cycle.

In human cell lines, Haspin depletion leads to a decrease

in H3T3 phosphorylation, defects in chromosome congres-

sion and a delay in exit from mitosis (Dai et al., 2005, 2006;

Markaki et al., 2009). In A. thaliana even in the context of an

organ, for AtHaspin RNAi down-regulated mitotic cells we

observed a reduced level of H3T3 phosphorylation but no

obvious defects of the mitotic cell cycle. In agreement with

this finding, flow cytometry analysis of young seedlings,

root tips and leaves from AtHaspin RNAi, Haspin

overexpression and WT plants revealed no significant

change in the proportion of 2C versus 4C nuclei, which

indicated that there was no severe alteration in the relative

duration of G1 and G2 (Figure S6). Further, a constant

number of chromosomes in plants with altered activity of

AtHaspin was confirmed by FISH in nuclei of differentiated

cells using a centromere-specific probe (Figure 8b). Taken

together these data suggest that a partial alteration of

AtHaspin transcripts is not sufficient to interfere with the

mitotic cell division in plants or that the response of Haspin

depletion in a cell culture is more pronounced that in the

context of a multicellular organ. Nevertheless, complete

depletion of AtHaspin showed cytokinesis defects during

early embryogenesis (Figure 4) that could explain the

essential function of AtHaspin for cell cycle progression in

plants. Moreover, in agreement with the previously made

observation that in plants H3T3 phosphorylation correlates

with the process of chromatin condensation (Houben et al.,

2007; Caperta et al., 2008), a slight reduction of chromosome

condensation was observed in AtHaspin-RNAi interphase

nuclei. This result confirms prior suggestions that the

distribution and function of phosphorylation on H3 of

mitotic chromosomes is almost reversed in plants compared

with mammals (Houben et al., 2007).

AtHaspin affects plant development as early as the

first embryonic cell division

The effect of Haspin on the development of a complex

organism was studied. The most striking effect of AtHaspin

on plant development was observed in young embryos.

Analysis of corresponding has1-3 mutant plants revealed

that the absence of AtHaspin activity results in embryo

lethality. The earliest stage at which we observed abnor-

malities in homozygous has1-3 mutant embryos was the

2-cell stage. In contrast with the WT, in which the apical cell

and the basal cell are in line with each other, in mutant

embryos the plane of the first cell division is skewed. This

observation suggests that the AtHaspin gene, like GNOM

(Mayer et al., 1993), is required before the asymmetric divi-

sion of the zygote. Subsequently, the pattern of further

divisions is changed in hypophysis and suspensor cells.

How AtHaspin influences the asymmetry of divisions of

the zygote is unknown. But interestingly, in mammals,

Haspin can be found at centrosomes in mitotic cells (Dai

et al., 2005), and Haspin RNAi leads to the emergence of

multiple acentriolar centrosome-like foci during mitosis (Dai

et al., 2009). Although plant cells have no centrosomes, it is

possible that Haspin in plants also interacts with compo-

nents of the cytoskeleton. As a consequence, AtHaspin

activity might influence the process of cytokinesis and the

symmetry of the cell division.

Phenotypical and transcriptional analysis of AtHaspin

mutants suggest an interplay between AtHaspin and auxin

transport

The observed growth phenotypes in AtHaspin mutants are

not explainable by a mitotic function of AtHapsin only.

Strong transcription activity of AtHaspin was observed in

proliferating tissues such as shoot apical meristem with

actively dividing cells, but only weakly, or not at all, in

differentiated tissues except the vasculature. Similar

transcription behavior has already been reported for genes

involved in the auxin transport pathway, such as PIN1

(Galweiler et al., 1998) and PID (Christensen et al., 2000), as

well as recycling of auxin transport component such as

GNOM which influence auxin transport by regulating the

endosomal recycling of PIN1 to the plasma membrane via

vesicle trafficking (Geldner et al., 2001, 2003).

We observed, as demonstrated for auxin transporter

mutants, an aberrant vascular patterning, reduced fertility

and lateral organs outgrowth (reviewed in Friml and

Palme, 2002). The observed abnormal cell division in early

embryogenesis resembled and was quite similar to the

GNOM gene mutation (Mayer et al., 1993; Geldner et al.,

2004). Alternatively, the polar localization and recycling of

PIN1 is mediated by GNOM (Geldner et al., 2004); there-

fore it is reasonable to hypothesize that these genes, which

are critical genes during embryogenesis, might be act with

AtHaspin in the same pathway. Furthermore, we could

show that the transcription levels of genes involved in

auxin transport pathway were reduced in both AtHaspin

RNAi and overexpressed mutants (Figure S4). As alteration

of these auxin-related genes are functioning in the same

biological pathway, this situation may explain similar

pleiotropic phenotype that have been observed in both

overexpressed and RNAi mutants. In addition these data

may indicate a possible direct or indirect interplay of

AtHaspin with those genes. Therefore, the potential func-

tion of AtHaspin in auxin transport pathway may explain

the observed expression of AtHaspin in the vascular

tissues. The challenge ahead is to define the precise

function, physiological regulation and signaling networks

of Haspin in plants.



EXPERIMENTAL PROCEDURES

Plant material and growth condition

Arabidopsis thaliana, ecotype ‘Columbia-0’ (Col-0) WT plants wereused in most of the studies. In addition, four different Arabidopsislines (SALK_019798, GABI435H08, GABI082D07, and GABI858F01)with T-DNA insertions in the AtHaspin gene obtained from theArabidopsis Biological Resource Center (ABRC) (http://www.Arabidopsis.org/abrc) were used. Plants were first grown under and 8-hphotoperiod, 22�C/18�C day/night temperature in controlled envi-ronment growth chambers. After 4 weeks plants were transferred tolong day conditions (16-h photoperiod per day).

Full-length cDNA and promoter isolation and cloning pro-

cedure

Arabidopsis thaliana cDNA synthesized by RevertAid H Minus firststrand cDNA synthesis kit (Fermentas, http://www.fermentas.com),was used for a PCR reaction to amplify the 1800-bp long codingregion of AtHaspin (primers AtHaspin-GateF and AtHaspin-GateR,listed Table S2). To isolate the putative promoter region, genomicDNA was used to amplify 1567 bp (primers AtHaspin-1567 pro F andAtHaspin-pro R, listed in Table S2) and 705 bp (primers AtHaspin-pro R and AtHaspin-705 pro F, listed in Table S2) upstream of thestart codon of AtHaspin using the proofreading Expand LongTemplate PCR enzyme kit (Roche, http://www.roche.com).

The Gateway pENTR Directional TOPO cloning kit (Invitrogen,http://www.invitrogen.com) was used to clone the blunt-ended PCRproduct into a pENTR/D-TOPO vector. After the TOPO cloningreaction, the pENTR TOPO construct was transformed to Escher-ichia coli strain DH5a (Stratagene, http://www.stratagene.com)by electroporation. The different destination (pEarleyGate100,pMDC107 and pMDC162 (Curtis and Grossniklaus, 2003; Earleyet al., 2006) vectors used in these experiments were obtained fromABRC or Invitrogen. Both pENTR destination vectors and pEarley-Gate vectors contained the same bacterial selection marker, namelykanamycin resistance. Therefore, to increase the efficiency ofrecombination, the entry vectors were linearized by MluI digestion.The LR Gateway recombination reaction was performed betweenentry and destination vector to recombine the sequence of interestinto the destination vector (e.g. pEarleyGate vectors) using the LRclonase II reaction (Invitrogen) according to the manufacture’sinstruction. The product of the LR reaction was transformed intoE. coli (DH5a strain) and plated on selective LB media containing50 mg L)1 kanamycin.

To reduce AtHaspin gene activity via RNAi, a hairpin RNAconstruct that contained a 400 bp AtHaspin highly specificsequence tag (GST), constructed in the pAGRIKOLA vector, wasobtained from NASC (The European Arabidopsis Stock Centre). Forplant transformation the Agrobacterium binary vectorGV3101::pSOUP (Hellens et al., 2000) was used. Transformation ofA. thaliana plants was performed by the floral dip method (Cloughand Bent, 1998).

Histology

For localization of promoter AtHaspin::GUS signals plant organs werefixed for 30 min in 90% acetone at room temperature. Organs werethen washed three times for 5 min each in washing buffer [0.1 M

sodium phosphate buffer, pH 7.0, 0.1% Triton X-100, 1 mM

K3Fe(CN)6, 1 mM K4Fe(CN)6] on ice. Subsequently, staining buffer[0.1 M sodium phosphate buffer, pH 7.0, 0.1% Triton X-100, 1 mM

K3Fe(CN)6, 1 mM K4Fe(CN)6, 1mM X-GLUC] was infiltrated by avacuum (approximately 15 min) and the samples were then incu-

bated at 37�C, overnight. GUS-stained organs were washed threetimes with 70% ethanol to remove chlorophyll.

For whole-mount embryo analysis siliques were opened longitu-dinally with a hypodermic needle, fixed in a mixture of ethanol andacetic acid (3:1) for 1 h and mounted in a drop of clearing solution(chloral hydrate: water: glycerol, 8w:3v:1v), as described by Weijerset al. (2001). Embryos were viewed with a Zeiss Axioplan IImicroscope equipped with differential interference contrast (DIC)optics.

Indirect immunofluorescence analysis

Organs of A. thaliana (root tips from 3-day-old seedlings or youngflower buds) were fixed for 20 min in freshly prepared 4% para-formaldehyde/phosphate-buffered saline (PBS) solution on ice,washed two times for 15 min each in PBS on ice, and digested at37�C for 25 min in a mixture of 2.5% cellulase ‘Onozuka R-10’ and2.5% pectolyase Y-23 dissolved in PBS. Organs were then washedfor 15 min in PBS. Squashed organs were subjected to immuno-staining as described (Houben et al., 2005). A rabbit polyclonalantibody against phospho-histone H3T3 (1:100, Cat. Nr. 07-424,Upstate) was used for immunodetection. Anti-rabbit-Cy3 was usedas secondary antibodies. 4¢,6-Diamidino-2-phenylindole (DAPI)antifade counterstained slides were analyzed with an OlympusBX61 microscope equipped with an ORCA-ER CCD camera.Deconvolution microscopy was employed for superior optical res-olution of globular structures. Thus each photograph was collectedas a sequential image along the Z-axis with approximately 11 slicesper specimen. All images were collected in gray scale andpseudocolored with Adobe Photoshop, and projections (maximumintensity) were done with the program Analysis (Soft ImagingSystem, http://www.soft-imaging.net/).

FISH analysis, image processing, and statistics

Preparation of nuclei, probe labeling, and FISH were as described(Schubert et al., 2008). FISH signals were analyzed with anepifluorescence microscope Axiophot (Zeiss, http://www.zeiss.de/)with a · 100x1.45 a-plan-fluar objective and a 3-chip color camera(DXC-950P; Sony, Tokyo, Japan). The microscope was integratedinto a Digital Optical 3D Microscope system (Confovis, http://www.confovis.com) to check signal separation/distances alongx-, y-, and z-axes. Images were captured separately for eachfluorochrome with appropriate excitation and emission filters. Theimages were merged with Adobe Photoshop 6.0 software (AdobeSystems, http://www.adobe.com). Chromosome territories (CTs)covering together more than 50% of the nucleus area wereregarded as dispersed (Figure 8d). FISH signals indicating posi-tional sister chromatid separation and chromatin dispersion werecompared against those of the Col-0 WT by the one-sided Fisher’sexact test.

Recombinant protein production

cDNA corresponding to the entire AtHaspin was cloned without thestop codon into vector pET101/D-TOPO (Invitrogen), using primersAtHaspin-without stop-F and AtHaspin-without stop-R (Table S2),and transformed to TOP10 competent cells (Invitrogen). Constructscontaining an empty vector, and mutation of lysine 310 to alanine,were produced by PCR-based mutagenesis (Phusion, Site-directedmutagenesis kit; Finnzymes, http://www.finnzymes.com/). All con-structs were confirmed by DNA sequencing. To generate C-6xHis-tag fusion peptides, pET101/D-TOPO plasmid AtHaspin, emptyvector and AtHaspin-mut constructs were transformed into E. coliBL21Star and inoculated into LB medium at 37�C. When culturesreached a density of 0.5–0.6 (OD 600) protein expression was



induced by adding 1 mM IPTG. After further incubation for 4–6 h at37�C, cells were collected by centrifugation. The recombinant pro-tein was purified using nickel-agarose columns (Ni-NTA spin kit;QIAGEN, http://www.qiagen.com) under native conditions accord-ing to the manufacture instruction.

In vitro kinase assay

Purified recombinant AtHaspin proteins were dialyzed against pro-tein storage buffer [20 mM HEPES (pH 7.4), 100 mM NaCl, 1 mM

DTT]. Approximately 4 lg AtHaspin was incubated for 60 min at30�C in the presence of kinase assay buffer [20 mM HEPES(pH 7.5),90 mM NaCl, 10 mM MgCl2, 6 mM MnCl2, 0.5 mM CaCl2, 100 lM

ATP], supplemented with 10 lCi [32P]ATP per sample. Twelvemicro gram core histone mix from calf thymus (Roche AppliedScience) and MBP were added as exogenous substrates per reac-tion. Reaction mixtures were resolved by 12% SDS-PAGE, stainedwith Coomassie Brilliant Blue, dried, and autoradiographed.Samples of the non-radioactive in vitro kinase assay were trans-ferred on to polyvinylidene fluoride membranes, and then themembranes were incubated with antibodies against histone H3[anti-total H3 (ab1791; Abcam, http://www.abcam.com/), as control],phosphorylated histone H3 at threonine 3 (07-424, Upstate), serine10 (06-570, Upstate), threonine 11 (Preuss et al., 2003) or serine 28(Goto et al., 1999).

ACKNOWLEDGEMENTS

We are grateful to Oda Weiss, Martina Kuhne and Katrin Kumke forexcellent technical assistance and Helmut Baumlein for discussionsand helpful comments on the manuscript. This work was supportedby the Land Sachsen-Anhalt (Network: structures and mechanismsof biological information processing) and the Deutsche Fors-chungsgemeinschaft (DFG, SFB 648).

SUPPORTING INFORMATION

Additional Supporting Information may be found in the onlineversion of this article:Figure S1. Model of AtHaspin gene structure and relative position ofthe T-DNA insertion mutations (has1-2, has1-2, has1-3 and has1-4).Figure S2. Multiple amino acid sequence alignment of haspin-likekinase domains of plants, human and mouse.Figure S3. Neighbor-joining tree of Haspin kinases calculated frompairwise median distances of the amino acid alignment of con-served parts of the sequences.Figure S4. Comparative transcription analysis of Auxin and devel-opmental related genes in AtHaspin mutant plants.Figure S5. Immunodetection of phosphorylated histone H3T3 inmitotic cells of 35S::AtHaspin overexpression plants.Figure S6. Relative proportion of 2C versus 4C nuclei in cotyledons,leaves and root tips of RNAi and/or Haspin overexpression(35S::Has) plants compared to wild-type.Table S1. Genotyping of progenies of the T-DNA heterozygousplants (Has/has1-3).Table S2. List of used primers.Please note: As a service to our authors and readers, this journalprovides supporting information supplied by the authors. Suchmaterials are peer-reviewed and may be re-organized for onlinedelivery, but are not copy-edited or typeset. Technical supportissues arising from supporting information (other than missingfiles) should be addressed to the authors.

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AtHaspin phosphorylates histone H3 at threonine 3 during mitosis and contributes to embryonic patterning in Arabidopsis