Research
FgKin1 kinase localizes to the septal pore and plays a role in
hyphal growth, ascospore germination, pathogenesis, and
localization of Tub1 beta-tubulins in Fusarium graminearum
Yongping Luo1*, Hongchang Zhang2*, Linlu Qi3, Shijie Zhang1, Xiaoying Zhou3, Yimei Zhang1 and Jin-Rong Xu1,3
1
State Key Laboratory of Crop Stress Biology for Arid Areas, College of Plant Protection, Northwest A&F University, Yangling, Shaanxi 712100, China; 2College of Life Sciences, Northwest
A&F University, Yangling, Shaanxi 712100, China; 3Department of Botany and Plant Pathology, Purdue University, West Lafayette, IN 47907-2054, USA
Summary
Author for correspondence:
Jin-Rong Xu
Tel: +1 765 496 6918
Email: jinrong@purdue.edu
Received: 8 May 2014
Accepted: 20 June 2014
New Phytologist (2014) 204: 943–954
doi: 10.1111/nph.12953
Key words: ascospore germination,
ascospore release, beta-tubulins, cytokinesis,
pathogenesis, septation.
The Kin1/Par-1/MARK kinases regulate various cellular processes in eukaryotic organisms.
Kin1 orthologs are well conserved in fungal pathogens but none of them have been functionally characterized. Here, we show that KIN1 is important for pathogenesis and growth in two
phytopathogenic fungi and that FgKin1 regulates ascospore germination and the localization
of Tub1 b-tubulins in Fusarium graminearum.
The Fgkin1 mutant and putative FgKIN1S172A kinase dead (nonactivatable) transformants
were characterized for defects in plant infection, sexual and asexual reproduction, and stress
responses. The localization of FgKin1 and two b-tubulins were examined in the wild-type and
mutant backgrounds.
Deletion of FgKIN1 resulted in reduced virulence and defects in ascospore germination and
release. FgKin1 localized to the center of septal pores. FgKIN1 deletion had no effect on Tub2
microtubules but disrupted Tub1 localization. In the mutant, Tub1 appeared to be enriched in
the nucleolus. In Magnaporthe oryzae, MoKin1 has similar functions in growth and infection
and it also localizes to septal pores. The S172A mutation had no effect on the localization and
function of FgKIN1 during sexual reproduction.
These results indicate that FgKIN1 has kinase-dependent and independent functions and it
specifically regulates Tub1 b-tubulins. FgKin1 plays a critical role in ascospore discharge, germination, and plant infection.
Introduction
The filamentous ascomycete Fusarium graminearum is one of the
causal agents of Fusarium head blight (FHB) of wheat and barley
(Bai & Shaner, 2004; Goswami & Kistler, 2004). It is also one of
the pathogens causing stalk and ear rots of maize. Unlike many
other plant pathogenic fungi, F. graminearum uses ascospores as
the primary inoculum to infect wheat or barley heads. The pathogen overwinters in infected plant tissues and produces perithecia
on plant debris. Ascospores are forcibly discharged from mature
perithecia (Trail et al., 2002; Trail, 2007) to infect wheat and
barley heads that are susceptible from anthesis to the dough stage
(Bai & Shaner, 2004). Asexual spores produced by this pathogen
on diseased plants are primarily for disease spreading. Under
favorable environmental conditions, F. graminearum can cause
severe yield losses and it produces harmful mycotoxins, such as
deoxynivalenol (DON) and zearalenone, in infected plant tissues.
As an inhibitor of protein synthesis in eukaryotic organisms,
DON is also an important virulence factor during plant
infection (Proctor et al., 1995; Bai et al., 2002). Mutants
*These authors contributed equally to this work.
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defective in DON production are defective in spreading from the
initial infection site via the rachis to the rest of wheat or barley
heads.
Because ascospores are the primary inoculum, sexual reproduction plays a critical role in the infection cycle of F. graminearum,
which is a homothallic fungus that contains two linked mating
type idiomorphs and four pheromone and pheromone receptor
genes (Kim et al., 2008; Lee et al., 2008; Zheng et al., 2013). In
addition to these well-conserved mating-related genes, a number
of genes with various biological functions are known to be important for sexual production in F. graminearum, including components of three well conserved mitogen-activated protein kinase
pathways and a number of transcription factor genes (Hou et al.,
2002; Jenczmionka et al., 2003; Urban et al., 2003; Son et al.,
2011; C. Wang et al., 2011; Nguyen et al., 2012). Whereas
mutants like the mgv1 and Gpmk1 deletion mutants were female
sterile and failed to form perithecia, some mutants, such as the
FgstuA, roa, zif1 and Gzrum1 mutants, were defective in ascospore formation or produced ascospores with abnormal morphology (Min et al., 2010; Kim et al., 2011; Lysoe et al., 2011; Y.
Wang et al., 2011). In addition, several genes, such as GEA1,
MID1, and CCH1, are known to be important for forcible
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discharge of ascospores in F. graminearum (Hallen & Trail,
2008; Cavinder et al., 2011; Son et al., 2013).
In an earlier study to functionally characterize the kinome of
F. graminearum (C. Wang et al., 2011), 26 protein kinase genes
were found to be important for ascospore production or release.
One of them is the FgKIN1 gene (FGSG_09274), which is orthologous to kin1 of Schizosaccharomyces pombe and KIN1/KIN2
of Saccharomyces cerevisiae. Kin1 orthologs belong to the family
of microtubule affinity-regulating protein kinases (MARKs),w
hich are involved in cellular polarization, cell cycle regulation, cell
migration and differentiation, cell signaling, and other biological
functions (Drewes et al., 1998; Tassan & Le Goff, 2004). In
addition to Kin1 in the yeasts, well studied members of the
MARK family include pEg3 of Xenopus and partitioning-defective 1 (PAR-1) of Caenorhabditis elegans and Drosophila.
Humans have four MARK isoforms, and defects in the tau kinase
have been associated with Alzheimer’s disease (Tassan & Le Goff,
2004). In S. cerevisiae, Kin1 and Kin2 are two paralogous MARK
proteins that interact with components of the secretary machinery and localize to the cytoplasmic face of the cytoplasm membrane (Tibbetts et al., 1994; Elbert et al., 2005). However, the
kin1 and kin2 mutants, even the kin1 kin2 double mutant, had
no obvious growth defects (Elbert et al., 2005). In S. pombe,
KIN1 is involved in morphogenesis, bipolar growth, intracellular
organization, and cytokinesis (Drewes & Nurse, 2003; La Carbona & Le Goff, 2006; Cadou et al., 2010). The kin1 deletion
mutant is defective in cell wall structure and cell morphology. In
Cryptococcus neoformans, the KIN1 ortholog was identified as a
virulence factor in infection assays with C. elegans (Mylonakis
et al., 2004). The kin1 disruption mutant is significantly reduced
in virulence but had no obvious morphological defects.
Although the MARK genes are well conserved, none of them
have been functionally studied in plant pathogenic fungi or filamentous ascomycetes. In this study, we characterized the role of
FgKIN1 in growth and development in F. graminearum and its
ortholog in the rice blast fungus Magnaporthe oryzae, a model for
studying fungal–plant interaction (Ebbole, 2007). The Fgkin1
and Mokin1 mutants were reduced in growth rate, conidiation,
and virulence. Both FgKin1 and MoKin1 localized to the center
of septal pores in living cells, although they are not essential for
hyphal tip growth and septum formation. In F. graminearum,
FgKin1 regulates the localization of Tub1, but not Tub2, b-tubulins to the microtubules. The FgKin1 protein has both kinasedependent and -independent functions but its localization to the
septal pore is independent of kinase activities. In addition,
FgKIN1 plays a critical role in conidiogenesis, pathogenesis, autoinhibition of ascospore germination, and ascospore release.
Materials and Methods
on carrot agar plates were assayed as previously described (Y.
Wang et al., 2011). To assay for defects in stress responses,
0.01% sodium dodecyl sulfate (SDS), 0.03% H2O2, 0.7 M
NaCl, 600 lg ml 1 Congo red, or 300 lg ml 1 Calcofluor was
added to PDA (Wang et al., 2012). Ascospore discharge was
assayed as described (Cavinder et al., 2012). Protoplasts were
prepared from 12 h germlings and used for polyethylene glycol
(PEG)-mediated transformation (Zhou et al., 2011). Hygromycin B (Roche) and geneticin (MP Biochemicals, Santa Ana, CA,
USA) were added to final concentrations of 300 lg ml 1 and
400 lg ml 1, respectively, in the regeneration medium.
Generation of the FgKIN1-GFP, FgKIN1S172A-GFP and H1GFP transformants
To generate the FgKIN1-GFP construct by gap repair, the entire
FgKIN1 gene, including its promoter region, was amplified with
primers KIN1-CM-F and KIN1-CM-R (Supporting Information, Table S1) and transformed with XhoI-digested pFL2 (Zhou
et al., 2011) into yeast strain XK1-25 (Bruno et al., 2004). The
resulting FgKIN1–GFP fusion construct carrying the geneticinresistant marker was transformed into the Fgkin1 mutant K5.
The resulting transformants were screened by PCR and confirmed by examination for green fluorescent protein (GFP) signals. The same gap repair approach was used to generate the
FgKIN1S172A–GFP construct by amplifying FgKIN1 with overlapping PCR using primers KD1 and KD2 (Table S1). Transformants of mutant K5 expressing the FgKIN1S172A–GFP construct
were identified by PCR and examination for GFP signals using
epifluorescence microscopy.
The RP27 promoter sequence was amplified with primers
RP27-F and RP27-R (Table S1) from vector pFL2 (Wang et al.,
2012) and cloned between the NotI and XbaI sites on pMF280
(Freitag et al., 2004). The resulting PRP27-H1–GFP construct
was confirmed by sequencing analysis and cotransformed into
PH-1 and the Fgkin1 mutant K5 with the geneticin-resistant vector pFL7 (Zhou et al., 2011). Transformants expressing the H1–
GFP construct were identified by PCR and confirmed by examination for GFP signals in the nucleus.
DAPI and Calcofluor staining
Freshly harvested conidia and hyphae were first fixed with 3.7%
formaldehyde and 0.2% Triton X-100 in 50 mM PBS buffer
(pH 7.0) for 30 min. After staining with 20 lg ml 1 Calcofluor
and 20 lg ml 1 4,6-diamidino-2-phenylindole (DAPI), samples
were examined with an Olympus BX53 fluorescence microscope
or an Olympus FV1000 confocal microscope. Perithecia were
cracked open to release asci and ascospores before DAPI and
Calcofluor staining.
Strains of F. graminearum and culture conditions
The wild-type strain PH-1 (Cuomo et al., 2007) and all the
transformants generated in this study were routinely cultured at
25°C on potato dextrose agar (PDA) or complete medium (Hou
et al., 2002). Growth rate, conidiation, and sexual reproduction
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Plant infection and DON assays
Flowering wheat heads of cv XiaoYan 22 or Norm were dropinoculated with 10 ll of conidium suspensions (2.0 9 105 conidia ml 1) as previously described (Gale et al., 2007). Scab
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symptoms were examined at 14 d postinoculation (dpi). At least
15 infected wheat heads were examined in each replicate to estimate the disease index for individual strains. DON production in
infected kernels was assayed as previously described (Bluhm et al.,
2007). For corn stalk rot assays, adult plants of cv Pioneer 2375
were inoculated with toothpicks soaked in conidium suspensions
for 1 min as previously described (Seong et al., 2006). Stalk rot
symptoms were examined at 14 dpi.
Generating the TUB1–GFP and TUB2–GFP transformants
The TUB1 gene was amplified with primers Tub1-eGFP-F and
Tub1-eGFP-R (Table S1) and cloned into pFL2 by the yeast gap
repair approach (Zhou et al., 2011). The same approach was used
to generate the TUB2–GFP fusion construct. The resulting
TUB1– and TUB2–GFP constructs were confirmed by sequencing analysis and transformed into protoplasts of PH-1 and the
Fgkin1 mutant K5. Transformants expressing the TUB1–GFP or
TUB2–GFP construct were identified by PCR and microscopic
examination for GFP signals.
Generation of transformants expressing the TUB2–,
TUB3–, and FgKIN1S172A–mCherry constructs and transformants
We first replaced GFP on pFL2 (Zhou & Xu, 2011) with
mCherry amplified from pE3279 to generate the mCherry vector
pMF36. The TUB2 gene was then cloned into pMF36 by yeast
gap repair (Bruno et al., 2004; Zhou & Xu, 2011) to generate the
TUB2–mCherry fusion. The TUB2–mCherry fusion construct
was cotransformed with FgKIN1–GFP into protoplasts of PH-1.
Transformants expressing TUB2–mCherry and FgKIN1–GFP
fusion constructs were identified by PCR and microscopic examinations for mCherry and GFP signals.
The c-tubulin gene TUB3 (FGSG_09993)–mCherry and
FgKIN1S172A–mCherry fusion constructs were generated with a
similar approach and confirmed by sequencing analysis. They
were cotransformed with the TUB1–GFP construct into protoplasts of the Fgkin1 mutant K5. All the resulting TUB1–GFP
TUB2–mCherry and TUB1–GFP FgKIN1S172A–mCherry transformants were verified by PCR and assayed for GFP or mCherry
signals.
Generation of the Mokin1 deletion mutant and MoKIN1–
GFP transformant in M. oryzae
The Ku80 strain (Villalba et al., 2008) and its transformants were
cultured on oatmeal agar for conidiation as described by Zhou
et al. (2012). The MoKIN1 deletion construct was generated by
the ligation-PCR approach. Protoplast preparation and transformation were performed as previously described (Ding et al.,
2010; Zhou et al., 2012). Putative Mokin1 deletion mutants were
identified by PCR and confirmed by Southern blot analysis. The
MoKIN1–GFP construct was generated by cloning the MoKIN1
gene into pDL2 by gap repair (Zhou et al., 2011). Appressorium
formation and plant infection were assayed with conidia
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harvested from 10-d-old oatmeal agar cultures as previously
described (Ding et al., 2010).
Results
The FgKIN1 MARK gene is not essential for growth but
important for conidiogenesis
Kin1 orthologs are well conserved in fungi. Like most fungal
MARK kinases, FgKin1 contains an N-terminal kinase domain and
a C-terminal kinase-associated domain 1 (KA1) domain (Tochio
et al., 2006; Moravcevic et al., 2010). Some members of the animal
MARK family, such as Par-1 and human MARK1-4, also have an
ubiquitin-associated domain, which is absent in fungal Kin1 proteins (Fig. S1). The Fgkin1 deletion mutants were identified in a
previous study of systemic characterization of protein kinase genes
in F. graminearum (C. Wang et al., 2011). In this study, two
Fgkin1 mutants, K3 and K5 (Table 1), were confirmed by Southern
blot analysis (Fig. S2). Mutants K3 and K5 had the same phenotype
although only data with K5 are described in the following.
The Fgkin1 mutant was reduced c. 19% in growth rate
(Table 2) and formed colonies with short, dense aerial hyphae
(Fig. 1a). However, it was reduced over 70% in conidiation
(Table 2). Conidia produced by the Fgkin1 mutant were shorter
and had fewer septa than those of the wild-type (Fig. 1b). When
stained with Calcofluor, 86.8 4.8% of the Fgkin1 conidia had
only three septa, but 91.2 4.6% of the wild-type conidia had
over four septa. The middle compartments of mutant conidia
were longer than those of PH-1 conidia. The Fgkin1 mutant also
had irregular compartment length in hyphae cultured in yeast
extract peptone dextrose (YEPD; Fig. 1c). These results indicated
that the Fgkin1 mutant was defective in septation during vegetative growth and asexual reproduction in F. graminearum.
FgKIN1 is important for virulence and tolerance to hyperosmotic and cell wall stresses
In infection assays with flowering wheat heads, the Fgkin1 mutant
caused typical scab symptoms in the inoculated wheat kernels
(Fig. 2a) and was able to spread to nearby spikelets (Table 2).
However, it had c. 50% reduction in virulence compared with
PH-1 (Fig. 2a; Table 2). The average disease indexes of the Fgkin1
mutant and PH-1 were 5 and 13, respectively (Table 2). In infection assays with corn stalks, the Fgkin1 mutant was also significantly reduced in virulence (Fig. 2b; Table S2). Therefore, FgKIN1
is important for full virulence in wheat and corn infection.
Because DON is an important virulence factor in
F. graminearum, we assayed DON production in infested wheat
kernels. No significant difference was observed between the wildtype and Fgkin1 mutant strains (Table 2). We also assayed defects
of the Fgkin1 mutant in response to different stresses and found
that it was normal in response to SDS or H2O2 but had increased
sensitivities to 0.7 M NaCl, 300 lg ml 1 Calcofluor, or
600 lg ml 1 Congo red (Fig. S3). These results indicate that
FgKIN1 may be important for tolerance to cell wall and hyperosmotic stresses.
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Table 1 The wild-type and mutant strains used in this study
Strains
Brief descriptions
Fusarium graminearum strains
PH-1
Wild-type
K3
Fgkin1 deletion mutant of PH-1
K5
Fgkin1 deletion mutant of PH-1
Fgtub1
tub1 deletion mutant
Fgkin1/FgKIN1–GFP transformant of K51
Fgkin1/FgKIN1–GFP transformant of K5
Transformant of PH-1 expressing H1–GFP
Transformant of K5 expressing H1–GFP
Transformant of K5 expressing H1–GFP
Transformant of K5 expressing H1–GFP
Transformant of PH-1 expressing
TUB1–GFP
T1-P11
Transformant of PH-1 expressing
TUB1–GFP
T1-K2
Transformant of K5 expressing TUB1–GFP
T1-K12
Transformant of K5 expressing TUB1–GFP
T1-K14
Transformant of K5 expressing TUB1–GFP
T2-P3
Transformant of PH-1 expressing
TUB2–GFP
T2-P4
Transformant of PH-1 expressing
TUB2–GFP
T2-K3
Transformant of K5 expressing TUB2–GFP
T2-K4
Transformant of K5 expressing TUB2–GFP
KD3
Transformant of K5 expressing
FgKIN1S172A–GFP
KD4
Transformant of K5 expressing
FgKIN1S172A–GFP
SJ23
FgKIN1–GFP and TUB2–mCherry
transformant of PH-1
T1-cMK2 TUB1–GFP and TUB3–mCherry
transformant of K5
T1-cMK3 TUB1–GFP and TUB3–mCherry
transformant of K5
T1-cMK4 TUB1–GFP and TUB3–mCherry
transformant of K5
T1-KDM2 TUB1–GFP and FgKIN1S172A–mCherry
transformant of K5
T1-KDM3 TUB1-GFP and FgKIN1S172A-mCherry
transformant of K5
T1-KDM4 TUB1–GFP and FgKIN1S172A–mCherry
transformant of K5
Magnaporthe oryzae strains
Ku80
Wild-type
C17
C19
HP6
HK2
HK3
HK4
T1-P10
Kin1-1
Kin1-2
Kin1-C
Mokin1 deletion mutant of Ku80
Mokin1 deletion mutant of Ku80
Mokin1/MoKIN1–GFP
transformant of Kin1-2
Table 2 Growth rate, conidiation, virulence, and deoxynivalenol (DON)
production assays with Fusarium graminearum strains
Reference
Cuomo
et al. (2007)
C. Wang
et al. (2011)
C. Wang
et al. (2011)
Qiu
et al. (2012)
This study
This study
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Conidiation
(106 ml 1)2
Disease
index3
DON
(ppm)4
18.7 0.2a5
15.2 0.2b
18. 9 0.1a
1.2 0.1a
0.4 0.0b
1.2 0.1a
11.0 2.7a
5.4 1.5b
11.2 2.8a
1401.0 93.1
1120.7 79.3
NA
1
Growth rate was measured as the daily extension of colony radium.
Conidia produced in 5-d-old CMC (CM with carboxymethylcellulose)
cultures. Mean and standard deviation (mean SD) were calculated from
at least three independent measurements.
3
Diseased spikelets per wheat head examined at 14 d postinoculation
(dpi).
4
DON production in the inoculated wheat kernels collected at 14 dpi.
5
Means SD were calculated with results from three independent
experiments. Data were analyzed with the protected Fisher’s least
significant difference (LSD) test. Different letters mark significant
differences (P = 0.05). NA, not assayed.
2
wild-type perithecia 2 wk after fertilization, perithecia of the
Fgkin1 mutant rarely produced cirrhi (Fig. 3a). For rare cirrhi
formed on the top of Fgkin1 perithecia, they had only limited
extension beyond the initial ooze (Fig. 3a), suggesting that ascospore release is defective in the Fgkin1 mutant. To confirm this
observation, we assayed for ascospore discharge as previously
described (Cavinder et al., 2012). Whereas abundant ascospores
were forcibly discharged from wild-type perithecia after incubation for 16 h, no ascospore discharge was observed in the Fgkin1
mutant (Fig. 3b), even after prolonged incubation up to 44 h.
Therefore, FgKIN1 plays a critical role in ascospore discharge.
This study
This study
This study
This study
This study
This study
Villalba
et al. (2008)
This study
This study
This study
1
All the fusion constructs were integrated ectopically in the
F. graminearum or M. oryzae genome.
Ascospore release is blocked in the Fgkin1 mutant
Perithecia produced by the Fgkin1 mutant on selfing plates had
normal morphology and size (Fig. 3a). However, whereas yellowish cirrhi were produced by the majority (74.2 3.5%) of
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PH-1 (WT)
K5 (kin1)
C17
(kin1/KIN1)
Growth rate
(mm d 1)1
Germination of Fgkin1 ascospores inside perithecia
Although cirrhi were rarely produced, asci with normal ascospores were produced in perithecia by the Fgkin1 mutant (Fig. 4a).
However, most of the mutant ascospores had germinated inside
perithecia 2 wk after fertilization and germinated ascospores were
tangled together by their germ tubes (Fig. 4b). Ascospore germination was not observed in 2-wk-old perithecia formed by the
wild-type and Fgkin1 complemented transformant. Thus, we
conclude that FgKIN1 is important for autoinhibition of ascospore germination inside perithecia.
The Fgkin1 mutant still produced four-celled ascospores with
one nucleus in each compartment (Fig. 4c), suggesting that it had
no defects in nuclear division and septation during ascospore formation. Interestingly, germ tubes were produced from only one
end of the mutant ascospores that germinated inside perithecia
(Fig. 4b). However, when ungerminated ascospores of the Fgkin1
mutant were incubated in liquid complete medium (CM), germination from both ends was observed (Fig. 4d). Therefore, the
mechanism regulating ascospore germination inside perithecia
must be different from that in nutrient media. We also assayed
germination with conidia harvested from 2-wk-old carrot agar
cultures. Conidium germination was not observed in freshly harvested conidia, although they were normal in germination when
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(a)
(b)
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(c)
Fig. 2 Defects in plant infection of the Fgkin1 mutant in Fusarium
graminearum. (a) Flowering wheat heads were drop-inoculated with
conidia of the wild-type (PH-1), Fgkin1 mutant (K5), and Fgkin1/FgKIN1
complemented transformant (C17). Black dots marked the inoculated
spikelets. Photographs were taken at 14 d postinoculation (dpi). (b) Corn
stalks were inoculated with conidia of PH-1, K5 and C17. Stalk rot
symptoms were observed at 14 dpi.
(a)
Fig. 1 Growth, conidium morphology, and septation defects of the Fgkin1
mutant in Fusarium graminearum. (a) Three-day-old complete medium
(CM) cultures of the wild-type (PH-1), Fgkin1 mutant (K5), and Fgkin1/
FgKIN1 complemented transformant (C17). (b) Conidia of the same set of
strains stained with Calcofluor and 4,6-diamidino-2-phenylindole (DAPI)
were observed by differential interference contrast (DIC, left) and
fluorescence (UV) microscopy. (c) Hyphae of transformants of PH-1 (HP6)
and K5 (HK2) expressing the H1–GFP construct and stained with
Calcofluor. Bars, 10 lm.
(b)
incubated in YEPD (Fig. S4), indicating that FgKIN1 must have
an ascospore-specific role for germination in F. graminearum.
Deletion of FgKIN1 affects the localization of Tub1 but not
Tub2 b-tubulins in F. graminearum
Microtubule affinity-regulating protein kinases phosphorylate
microtubule-associated proteins (MAPs) at the tubulin binding
sites to induce their detachment from microtubules (Tassan & Le
Goff, 2004). To determine the role of FgKIN1 in microtubule
organization, we generated the TUB1 (FGSG_09530)–GFP
fusion construct and transformed it into PH-1 and the Fgkin1
mutant K5. As expected, GFP signals were observed in the microtubules in hyphae of the TUB1–GFP transformant of PH-1
(Fig. 5a). Surprisingly, in the Fgkin1/TUB1–GFP transformant
T1-K2 (Table 1), GFP signals were not localized to the microtubule cytoskeleton in hyphae (Fig. 5a), conidia (Fig. S5), ascospores, and germ tubes (Fig. S6). By contrast, it appeared that
Tub1–GFP proteins were aggregated in the nucleus (Fig. 5a). To
determine whether Tub1 localized to the nucleus, nuclei were
stained with DAPI in the TUB1–GFP transformant. We found
that Tub1–GFP fusion proteins localized to the nucleus but were
not evenly distributed (Fig. 5c). DAPI staining was weaker in the
region where strong Tub1–GFP signals were observed (Fig. 5c),
suggesting that deletion of FgKin1 might result in the
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Fig. 3 The Fgkin1 mutant in Fusarium graminearum was defective in
ascospore release. (a) Three-week-old mating cultures of the wild-type
(PH-1), Fgkin1 mutant (K5), and Fgkin1/FgKIN1 complemented
transformant (C17). The Fgkin1 mutant rarely formed cirrhi that had
limited extension in comparison with those of the wild-type. Cirrhi (masses
of ascospores) that have oozed out from the ostioles of perithecia are
marked with arrows. Bar, 200 lm. (b) Ascospore discharge was assayed
with 1-wk-old perithecia of PH-1, K5, and C17. Accumulation of
ascospores released from perithecia (visible as whitish masses) was
examined after incubation for 16 h. The Fgkin1 mutant had no visible
ascospore discharge.
localization of Tub1 to the nucleolus, which is not well stained
with DAPI (Banuett & Herskowitz, 2002; Fox et al., 2002).
Unlike most filamentous fungi, F. graminearum contains two
b-tubulin genes. TUB2 (FGSG_06611) plays a much more critical role than TUB1 in hyphal growth and fungicide resistance
(Chen et al., 2009; Qiu et al., 2012). Because transformant T1K2 was similar to the Fgkin1 mutant in growth and reproduction,
the localization of TUB2 to microtubules should not be affected
by FgKIN1 deletion. To test this hypothesis, we also generated
the TUB2–GFP construct and transformed it into PH-1 and
mutant K5. In the resulting transformants (Table 1), Tub2–GFP
mainly localized to the microtubules in hyphae (Fig. 5b). No differences in the localization of Tub2–GFP were observed between
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(a)
(a)
(b)
(b)
A
B
C
D
(c)
(c)
(d)
(d)
Fig. 4 Defects in ascospore germination of the Fgkin1 mutant in Fusarium
graminearum. (a) Asci with ascospores in perithecia of PH-1, Fgkin1
mutant K5, and complemented transformant C17. (b) Ascospores of the
Fgkin1 mutant (B, C) germinated inside 10-d-old perithecia. Ascospore
swelling but not germination was observed in PH-1 (A) and C17 (D). (c)
Ascospores of PH-1 and K5 were stained with Calcofluor and 4,6diamidino-2-phenylindole (DAPI). The Fgkin1 mutant still produced fourcelled ascospores with one nucleus in each cell compartment that
germinated inside perithecia. (d) Ungerminated ascospores of PH-1, K5,
and C17 were incubated in complete medium (CM) at 25°C for 12 h.
Bars, 20 lm.
the wild-type and Fgkin1 mutant transformants. Therefore,
FgKin1 is dispensable for the formation of Tub2 microtubules.
The Kin1 kinase must play a specific role in regulating the localization or organization of Tub1 b-tubulins in F. graminearum.
Enrichment of Tub1 in the nucleus in the Fgkin1 mutant is
not related to the MTOC
Because the microtubule organizing center (MTOC) or spindle
pole body (SPB) is the structure next to the nucleus that consists
of b-tubulins, Tub1-GFP may be disorganized and aggregated
near the MTOC in the Fgkin1 deletion mutant. To test this
hypothesis, we generated the TUB3–mCherry construct and cotransformed it with TUB1–GFP into PH-1 and the Fgkin1
mutant K5. The TUB3 gene (FGSG_09993) encodes the
gamma-tubulin that is a marker for MTOCs (Ohta et al., 2012).
In the resulting TUB1–GFP and TUB3–mCherry transformants
(Table 1), Tub3–mCherry localized to the MTOC (Fig. 5d).
However, Tub1–GFP signals appeared to be enriched in an area
that is not related to the MTOC in the Fgkin1 mutant. Whereas
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Fig. 5 Deletion of FgKIN1 in Fusarium graminearum affected the
subcellular localization of Tub1 but not Tub2 b-tubulins. (a) TUB1–GFP
transformants of PH-1 and the Fgkin1 mutant K5 were examined by
differential interference contrast (DIC) and fluorescence microscopy. (b)
Hyphae of the transformants of PH-1 and K5 expressing the TUB2–GFP
construct. (c) The TUB1–GFP transformants of PH-1 and K5 were
examined by confocal microscopy after staining with 4,6-diamidino-2phenylindole (DAPI) and Calcofluor. (d) Hyphae of the transformant T1–
cMK2 of the Fgkin1 mutant K5 expressing the TUB1–GFP and TUB3–
mCherry fusion constructs were assayed for the localization of Tub1–GFP
and Tub3–mCherry fusion proteins. Bars, 10 lm.
in some nuclei, Tub1–GFP proteins aggregated at the opposite
side of the MTOC, they were adjacent to each other in other
nuclei (Fig. 5d).
Localization of FgKin1 to the septal pore
For complementation assays, the FgKIN1–GFP fusion construct
was transformed into the Fgkin1 mutant strain K5. The resulting
complemented transformant strain C17 has all the phenotype
rescued. It was normal in growth (Fig. 1a), plant infection
(Fig. 2), and sexual reduction (Fig. 3). When examined by fluorescence microscopy, GFP signals were mainly observed in the
center of septal pores in conidia and hyphae (Fig. 6a). Close
examination by confocal microscopy revealed that FgKIN1–GFP
localization was slightly off the septum plate to the tip side of
hyphae and conidia (Video S1). However, no GFP signals were
observed at the tips of germ tubes and vegetative hyphae (Fig. 6a,
panel C). Together with normal conidium germination and
hyphal tip growth in the Fgkin1 mutant, these data suggest that
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FgKin1 is not important for establishing or maintaining polarized growth in F. graminearum.
Because the Fgkin1 mutant was defective in ascospore discharge and germination, we also examined the expression and
subcellular localization of FgKIN1–GFP during sexual reproduction. Similar to its localization in conidia, FgKin1–GFP localized
to the center of septal pores in ascospores and germ tubes produced by ascospores (Fig. 6b).
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(a)
A
C
B
D
FgKin1 localizes to the septal pore after septation is
complete
To determine the timing of FgKin1 association with the septal
pore, we examined the localization of FgKin1–GFP during septum formation. In the FgKIN1–GFP transformant, GFP signals
were not observed at the septation sites during early constriction
stages (Fig. 7a). The localization of FgKin1–GFP to the septal
pore was observed only at late stages of septum formation or
when septation is complete and the septum is mature (Fig. 7a).
Although FgKin1 is not involved in initial steps of septum formation, it may be important to maintain the function of septal
pores in F. graminearum. To test this hypothesis, we cotransformed the FgKIN1–GFP and TUB2–mCherry fusion constructs
into PH-1. In the resulting transformant SJ23 (Table 1), FgKin1
localized to the septal pore area where microtubules aggregated
between two cell compartments (Fig. 7b). The localization of
FgKin1 to the center of septal pores was also observed at the septum that separated the intact hyphal compartment from the damaged one (Fig. 7c). Therefore, FgKin1 and its orthologs may be
associated with the septal pore for septum functions in filamentous ascomycetes.
Kinase activity is not essential for the localization and
function of FgKin1 during sexual reproduction
To determine whether the kinase activity is essential for FgKin1
function and localization, we generated the FgKIN1S172A–GFP
allele and transformed it into mutant K5. The S172 residue of
FgKin1 is equivalent to S212 of MARK2, which is essential for
the kinase activity (Timm et al., 2008). The resulting Fgkin1/
FgKIN1S172A–GFP transformants KD3 (Table 1) had similar
defects in growth rate, virulence, and conidium morphology
(Fig. 7a) to the original Fgkin1 mutant (Fig. 8a). However,
FgKIN1S172A–GFP fusion proteins still localized to the septal
pore in conidia and hyphae of transformant KD3 (Fig. 8a). These
results suggested that the kinase activity is essential for the
FgKin1 function but dispensable for its subcellular localization
during vegetative growth and asexual reproduction.
Interestingly, unlikely the Fgkin1 mutant, the Fgkin1/
FgKIN1S172A transformants produced cirrhi at 2 wk after fertilization. Forcible discharge of ascospores was also observed in the
Fgkin1/FgKIN1S172A–GFP transformants although at a reduced
level in comparison with PH-1 (Fig. 8b). Inside perithecia, most
of the ascospores were not germinated 2 wk after fertilization
(Fig. 8c), indicating that the kinase activity is not essential for the
function of FgKIN1 in ascospore germination and discharge.
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(b)
Fig. 6 Subcellular localization of the FgKin1–GFP fusion protein in
Fusarium graminearum. (a) Conidia (upper row) and 12 h germlings
(lower row) of the Fgkin1/FgKIN1–GFP transformant C17 were stained
with Calcofluor and examined by confocal microscopy. FgKin1 localized to
the center of septum pores in conidia (A and B) and germlings. Whereas
panels A and C were regular stacked images, the images in panels B and D
were tilted to show the central localization of FgKin1 to the septal pore. (b)
Ascospores (upper row) and germinated ascospores (lower row) of the
FgKIN1–GFP transformant C17 were examined by differential interference
contrast (DIC) and epifluorescence (GFP) microscopy. Arrows point to the
localization of FgKin1–GFP at the septal pore. Bars, 10 lm.
Therefore, the FgKin1 protein may have both kinase-dependent
and -independent activities in F. graminearum, and possibly other
filamentous fungi.
We also cotransformed the TUB1–GFP and FgKIN1S172A
constructs into the Fgkin1 mutant. GFP signals were mainly
observed in the nucleus (Fig. S7) in the resulting transformant
T1–KDM2 (Table 1), which was similar to what was observed
in the Fgkin1 TUB1–GFP transformant (Fig. 5a). Therefore,
Tub1 localization is not dependent on the kinase activity of
FgKin1.
MoKin1 has similar function and localization with FgKin1
Because Kin1 orthologs have not been characterized in other filamentous ascomycetes, we also identified and characterized the
MoKIN1 (MGG_01279) gene in the rice blast fungus M. oryzae.
Similar to the Fgkin1 mutant, the Mokin1 deletion mutant was
reduced in growth rate (Table S3) and virulence in infection
assays with barley leaves (Fig. 9a; Table S3). However, deletion
of MoKIN1 had no effect on appressorium formation and penetration. In transformants expressing the MoKIN1–GFP construct, GFP signals also localized to the septal pore (Fig. 9b).
These results indicate that the localization of Kin1 orthologs and
their functions in hyphal growth and pathogenesis may be conserved in plant pathogenic fungi.
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(a)
(a)
(b)
(c)
(b)
Fig. 8 Localization and functions of the FgKin1S172A protein in Fusarium
graminearum. (a) GFP signals were observed in the septal pore in conidia
and hyphae of the Fgkin1/FgKIN1S172A–GFP transformant KD3. DIC,
differential interference contrast. (b) Ascospore discharge assays with
perithecia of PH-1 and transformant KD3. (c) Asci and ascospores
produced by transformant KD3. The close-up view on the right shows that
ascospores of KD3 had normal morphology. Bars, 10 lm.
(c)
Fig. 7 Association of FgKin1 with the septal pore during septum formation
in Fusarium graminearum. (a) A section of a hypha of the FgKIN1–GFP
transformant C17 was examined at the indicated time intervals by
differential interference contrast (DIC) and epifluorescence (GFP)
microscopy. The localization of FgKin1 to the septal pore was not observed
at the developing septum (marked with an arrow). GFP signals were
observed in the septal pore area only when the septum was mature. (b)
Conidia and hyphae of transformant SJ23 expressing the FgKIN1–GFP
with TUB2–mCherry fusion constructs. Colocalization of FgKin1 and Tub2
was observed at the septal pore site where microtubules aggregate
(marked with an arrow). (c) Localization of FgKin1–GFP to the plugged
septum (marked with an arrow) that separated the intact hyphal
compartment from the damaged part in transformant SJ23. The damaged
compartment lacked cytoplasm and Tub2–mCherry signals. Bars, 10 lm.
In appressorium formation assays with the MoKIN1–GFP
transformant, GFP signals were observed at the center of septal
pores in conidia at early stages (Fig. 9b). However, when appressoria were mature by 24 h, the localization of MoKin1–GFP to
the septal pore in conidia was no longer visible, although small
vesicles with GFP signals were observed in appressoria (Fig. 9b).
In M. oryzae, conidial compartments become dead and collapsed
when appressoria are mature (Veneault-Fourrey et al., 2006).
One septum was formed to delimit appressoria from the rest of
germ tubes and this septum is complete (no septal pore) for
appressorium turgor generation. In the MoKIN1–GFP
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transformant, GFP signals were not observed at the septum
delimiting melanized appressoria by 24 h (Fig. 9b), suggesting
that MoKin1 only localizes to the center of functional septal
pores in living cells.
Invasive hyphae formed by M. oryzae are known to be morphologically different from vegetative hyphae and they grow as
pseudohyphae (Zhou et al., 2012). We also examined the localization of MoKin1–GFP in invasive hyphae formed by the
MoKIN1–GFP transformant inside rice leaf sheath cells. GFP signals were observed at the center of the constriction sites that
delimit invasive hyphae into fragments (Fig. 9c), suggesting that
these constriction sites in invasive hyphae are structurally similar
to septa in vegetative hyphae.
TUB1 is also important for normal growth, conidiation,
conidiogenesis, and sexual reproduction
Because deletion of FgKIN1 disrupted the localization of Tub1,
we obtain the tub1 deletion mutant (Qiu et al., 2012). In comparison with PH-1, the tub1 mutant was reduced 70.4 3.0% in
growth rate (Fig. S8a) and 70.8 5.6% in conidiation. In addition, c. 95.8 1.5% of tub1 conidia had three or fewer septa
(Fig. S8b). These phenotypes of the tub1 mutant were similar to
those of the Fgkin1 mutant. Interestingly, the tub1 mutant still
produced few small perithecia that were sterile and contained no
asci or ascospores. We also noticed that colony morphology is
different between the Fgkin1 and tub1 mutants. These results
indicate that the defects of the tub1 mutant were more severe
than those of the Fgkin1 mutant.
Discussion
Kin1 kinases are members of the KIN1/Par-1/MARK family proteins that are involved in cell polarity and microtubule-based
transportation via phosphorylation of MAPs. Unlike most protein kinases, the C-terminal KA1 domain of MARKs is important for the autoinhibition of the kinase domain (Tochio et al.,
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(a)
(b)
(c)
Fig. 9 Defects of the Mokin1 mutant in pathogenesis and localization of
MoKin1–GFP in Magnaporthe oryzae. (a) Barley leaves inoculated with
Ku80, Mokin1 mutants Kin1-1 and Kin1-2, and Kin1-C. Inoculation with
gelatin solution was used as the control. (b) Conidia of the MoKIN1–GFP
transformant were examined for GFP signals after incubation on
hydrophobic coverslips for 0, 8 and 24 h. MoKin1 localized to the septal
pore at early stages but dispersed into small vesicles by 24 h. The cell wall
was stained with Calcofluor. Bar, 10 lm. (c) GFP signals were observed at
the center of the junction sites (arrows) between fragments of invasive
hyphae formed by the MoKIN1–GFP transformant in epidermal cells of
rice leaf sheaths.
2006; Moravcevic et al., 2010). In mammalian cells, the KA
domain can be divided into two regions. The N-terminal 50 aa
of KA1 is necessary for proper folding of the functional C-terminal 50 aa region with the ELKL motif. Interestingly, Kin1 kinases
from filamentous fungi and S. cerevisiae have an additional 50–
100 amino acids between these two KA1 regions. In comparison
with the well-conserved KA1 regions, fungal-specific sequences
vary more significantly and may be important for the function or
localization of Kin1 kinases in fungi.
The Fgkin1 and Mokin1 deletion mutants were reduced in vegetative growth. In S. pombe, the Kin1 kinase regulates cell extension and division, possibly by regulating microtubule density and
stability (Gladfelter & Berman, 2009; Galjart, 2010). In the
wild-type strain of F. graminearum, both Tub1 and Tub2 b-tubulins are associated with microtubules. Deletion of FgKIN1 had
no impact on Tub2 microtubules, but Tub1 became aggregated
in the nucleolus. In F. graminearum, TUB2 plays a more important role in vegetative growth than TUB1 (Chen et al., 2009; Qiu
et al., 2012). When the tub1 mutant (Chen et al., 2009; Qiu
et al., 2012) was compared with the Fgkin1 mutant, we found
that they were both reduced in growth rate and conidiation. Conidia produced by the tub1 mutant also had fewer septa and were
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smaller. Therefore, the defects of the Fgkin1 mutant in growth
and asexual reproduction may be directly related to the disruption of the formation of Tub1 microtubules.
Sexual reproduction plays a critical role in the disease cycle of
F. graminearum, but the Fgkin1 mutant was defective in ascospore discharge. In mature perithecia, mutant ascospores germinated and the resulting germ tubes tangled together, which may
be related to its defect in cirrhus formation and ascospore discharge. To our knowledge, it is not clear what mechanisms regulate autoinhibition of ascospore germination inside perithecia in
Sordariomycetes. In F. graminearum, FgKin1 may be directly
involved in suppressing ascospore germination before they are
released from perithecia. Whereas mutant ascospores were normal in germination when cultured in CM, they produced germ
tubes from one end inside perithecia, indicating that ascospore
germination may involve different regulatory mechanisms in cultures from those in perithecia. Because four-celled ascospores are
symmetrical in cellular structure, it is puzzling how
F. graminearum distinguishes two ends of ascospores in the
absence of FgKin1. Nevertheless, ascospores are arranged in the
order whereby one end is close to the operculum of asci. Interestingly, we found that the tub1 mutant rarely produced perithecia
and failed to form asci and ascospores. Although Tub2 is more
important for vegetative growth, it is dispensable for ascospore
formation and release, indicating that Tub1 plays a more critical
role in sexual reproduction in F. graminearum (Qiu et al., 2012).
Therefore, it remains possible that the defect of the Fgkin1
mutant in ascospore germination and release is also somehow
related to the dislocalization of Tub1.
In S. pombe, the kin1 mutant is delayed in septation and has
increased sensitivity to cell wall stressors (Cadou et al., 2010,
2013). In F. graminearum, the Fgkin1 mutant had irregular
septation in hyphae and increased sensitivity to Calcofluor and
Congo Red. It is possible that deletion of FgKIN1 adversely
affected cell wall integrity. In the Fgkin1 mutant, conidia had
fewer septa, but conidium germination was not affected. Interestingly, septation in ascospores appeared to be normal, indicating that the role of FgKIN1 in septation is different between
ascospores and conidia.
The Fgkin1 mutant was significantly reduced in virulence (c.
50%). One contributing factor to its reduced virulence is that the
Fgkin1 mutant was reduced in growth (19%). However, the
degree of reduction was more significant than that of reduced
growth rate, indicating that deletion of FgKIN1 may result in
additional defects related to infectious growth or overcoming
plant defense responses. Although the mutant was normal in
terms of DON production, it was reduced in septation and had
increased sensitivity to cell wall and hyperosmotic stresses. Our
data also suggested that FgKin1 plays a role in septal pore functions. In addition, the subcellular distribution and organization
of the Tub1 b-tubulins were affected by FgKIN1 deletion. In
C. neoformans, the Kin1 ortholog is also important for virulence
(Mylonakis et al., 2004). To date, Kin1 orthologs have not been
characterized in other plant pathogenic fungi. In this study, we
found that MoKIN1 is dispensable for appressorium formation
but important for full virulence in M. oryzae. Therefore, the role
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of Kin1 orthologs in pathogenesis may be conserved in fungal
pathogens.
In S. cerevisiae, Kin1 localizes to the cytoplasmic face of the
plasma membrane at the budding neck (Tibbetts et al., 1994;
Elbert et al., 2005). In S. pombe, Kin1 localizes to the membrane
of new mitotic cell ends or the active cell surface remodeling sites
(Drewes & Nurse, 2003; Cadou et al., 2010). However, in both
F. graminearum and M. oryzae, Kin1 localized to the center of
septal pores, suggesting that this well-conserved protein kinase
may be functionally related to the maintenance of cell or compartment partitioning. Localization of FgKin1 or MoKin1 to the
cytoplasm membrane was not observed, which may be related to
additional sequences present in the KA1 domain that is known to
target MARKs in mammalian cells to the cytoplasm side of the
negatively charged membrane. In dead conidium cells or empty
hyphal fragments, localization of MoKin1 to the septal pore was
not observed, indicating that the association of Kin1 with septal
pores is specific to functional septa in living cells. Although Kin1
appeared to be at the tip side of septa, we failed to observe the
localization of FgKin1– and MoKin1–GFP to hyphal tips. These
data, together with normal spore germination and hyphal growth
in the mutants, suggest that the Kin1 kinase is not essential for
establishing and maintaining polarized growth in filamentous
fungi.
Microtubules consist of heterodimers of a- and b-tubulin and
the b- and a-tubulin exposure ends are termed plus and minus
ends, respectively. In animal cells, the minus end localizes to the
MTOC in the middle of cell, while the plus end of microtubules
radiates towards epidermis (Galjart, 2010). In Drosophila, Par-1
mutation leads to the mislocalization of the plus end to the cell
center and an increase in microtubule density (Doerflinger et al.,
2003). In F. graminearum, localization of the Tub2 b-tubulin
that is important for growth, virulence, and benzimidazole fungicide resistance (Chen et al., 2009) was not affected by FgKIN1
deletion. However, deletion of FgKIN1 disrupted the organization and localization of Tub1 to microtubules and resulted in its
uneven accumulation in the nucleus. Although we originally
hypothesized that Tub1 may be overaggregated at the MTOC,
the area where Tub1–GFP proteins were enriched was different
from the localization of the Tub3 c-tubulins (Fig. 5d). In the
Fgkin1 mutant, Tub1 may be aggregated in the nucleolus,
because this region had faint DAPI staining and the nucleolus is
not well stained with DAP (Banuett & Herskowitz, 2002; Fox
et al., 2002). In mammalian cells, nuclear accumulation of soluble tubulins has been observed in tumor cells (Xu & Luduena,
2002; Akoumianaki et al., 2009).
The paralogous Tub1 and Tub2 proteins are highly similar to
each other. They share 76% identity and have variations throughout the entire protein (Fig. S9). It will be important to determine
the regions of Tub1 and Tub2 that are responsible for binding to
different MAP proteins. F. graminearum may have Tub1-specific
MAP proteins that are phosphorylated by FgKin1. Interestingly,
we noticed that various fungi belonging to different phyla contain
two b-tubulin genes. The Kin1 orthologs may be evolutionally
conserved in these fungi to differentially regulate the organization
of these two b-tubulins into cytoskeleton microtubules.
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In the FgKIN1S172A transformant, defects of the Fgkin1
mutant in growth, asexual reproduction, and Tub1 aggregation
were not rescued. However, expression of the putative kinasedead allele partially suppressed the ascospore germination and
discharge defects. Therefore, FgKin1 must have kinase-dependent and -independent functions in F. graminearum. It has been
well documented that some protein kinases, such as Kss1, possess
functions independent of kinase activity (Breitkreutz et al., 2001;
Sabbagh et al., 2001). However, kinase-independent activity has
not been reported in Kin1 orthologs. In F. graminearum, the
FgKin1S172A protein still localizes to the septal pore, indicating
that the subcellular localization of FgKin1 is independent of its
kinase activity. It is possible that certain FgKin1-interacting proteins are responsible for anchoring the wild-type or putative
kinase-dead proteins to the center of septal pores. Septum formation in filamentous ascomycetes is different from septation in
yeast cells and basidiomycetes. The subcellular localization of
Kin1 may be associated with microtubule organization and septum function in filamentous ascomycetes. Therefore, it will be
important to identify and characterize Kin1-interacting proteins
or MAPs phosphorylated by Kin1 in F. graminearum and other
fungi.
Acknowledgements
We thank Drs Huiqian Liu and Chenfang Wang at
Northwest A&F University for fruitful discussions. We also
thank Dr MingGuo Zhou at Nanjing Agricultural University for
providing the tub1 deletion mutant. This work was supported by
the National Major Project of Breeding for New Transgenic
Organisms (2012ZX08009003), the National Basic Research
Program of China (2013CB127703 and 2012CB114002), and
the Special Fund for Agroscientific Research in the Public
Interest (201303016-6).
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Supporting Information
Additional supporting information may be found in the online
version of this article.
Fig. S1 Fusarium graminearum Kin1 and its orthologs from other
fungi.
Fig. S2 The FgKIN1 gene replacement construct and deletion
mutants in Fusarium graminearum.
New Phytologist (2014) 204: 943–954
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Fig. S3 Assays for responses of the Fgkin1 mutant to different
stresses in Fusarium graminearum.
Fig. S4 Conidium germination assays of PH-1 and the Fgkin1
mutant in Fusarium graminearum.
Fig. S5 Localization of Tub1 and Tub2 in conidia of transformants of PH-1 and the Fgkin1 mutant K5 expressing the TUB1or TUB2-GFP fusion construct in Fusarium graminearum.
Fig. S6 Localization of Tub1 in germ tubes and ascospores of
transformant of the Fgkin1 mutant K5 expressing the TUB1GFP fusion construct in Fusarium graminearum.
Fig. S7 Localization of Tub1-GFP in the FgKIN1S172A transformant T1-KDM2 in Fusarium graminearum.
Fig. S8 Phenotypes of the Fgkin1 and tub1 mutants in Fusarium
graminearum.
Fig. S9 Alignment of the amino acid sequences of Tub1 and
Tub2 in Fusarium graminearum.
Table S1 PCR primers used in this study
Table S2 Defects in infection assays with corn silks and stalks of
the Fgkin1 mutant in Fusarium graminearum
Table S3 Defects in vegetative growth and plant infection of the
Mokin1 mutant in Magnaporthe oryzae
Video S1 Localization of FgKin1–GFP proteins to the center of
septal pores in conidia of the Fgkin1/FgKIN1–GFP transformant
C17 in Fusarium graminearum.
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