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Courtship Ritual of Male and Female Nuclei during Fertilization
in Neurospora crassa
Sylvain Brun,
a
Hsiao-Che Kuo,
c
Chris E. Jeffree,
c
Darren D. Thomson,
b
*Nick Read
b
,
c†
a
Laboratoire Interdisciplinaire des Energies de Demain, CNRS UMR 8236, Université de Paris, Paris, France
b
Manchester Fungal Infection Group, Division of Infection, Immunity, and Respiratory Medicine, University of Manchester, Manchester, United Kingdom
c
Fungal Cell Biology Group, Institute of Cell Biology, University of Edinburgh, Edinburgh, United Kingdom
ABSTRACT Sexual reproduction is a key process influencing the evolution and ad-
aptation of animals, plants, and many eukaryotic microorganisms, such as fungi.
However, the sequential cell biology of fertilization and the associated nuclear dy-
namics after plasmogamy are poorly understood in filamentous fungi. Using histone-
fluorescent parental isolates, we tracked male and female nuclei during fertilization
in the model ascomycete Neurospora crassa using live-cell imaging. This study unrav-
els the behavior of trichogyne resident female nuclei and the extraordinary manner
in which male nuclei migrate up the trichogyne to the protoperithecium. Our obser-
vations raise new fundamental questions about the modus operandi of nucleus
movements during sexual reproduction, male and female nuclear identity, guidance
of nuclei within the trichogyne and, unexpectedly, the avoidance of “polyspermy”in
fungi. The spatiotemporal dynamics of male nuclei within the trichogyne following
plasmogamy are also described, where the speed and the deformation of male
nuclei are of the most dramatic observed to date in a living organism.
IMPORTANCE Using live-cell fluorescence imaging, for the firsttimewehaveobserved
live male and female nuclei during sexual reproduction in the model fungus Neurospora
crassa.Thisstudyrevealsthespecific behavior of resident female nuclei within the tri-
chogyne (the female organ) after fertilization and the extraordinary manner in which
male nuclei migrate across the trichogyne toward their final destination, the protoperi-
thecium, where karyogamy takes place. Importantly, the speed and deformation of male
nuclei were found to be among the most dramatic ever observed in a living organism.
Furthermore, we observed that entry of male nuclei into protoperithecia may block the
entry of other male nuclei, suggesting that a process analogous to polyspermy avoid-
ance could exist in fungi. Our live-cell imaging approach opens new opportunities for
novel research on cell-signaling during sexual reproduction in fungi and, on a broader
scale, nuclear dynamics in eukaryotes.
KEYWORDS sexual reproduction, fertilization, nucleus, trichogyne, mating, live-cell
imaging, fungi, Neurospora crassa,filamentous fungi
Fungi are living microorganisms that considerably impact human life. These adaptive
eukaryotes have been shown to thrive in the damaged Chernobyl radioactive reactors
(1), space (2), and extreme temperatures (3). Their impressive growth capacity is used for
biotechnology and industrial purposes (4, 5). Fungal plant pathogens are responsible for
the loss of billions of dollars in food security due to crop losses worldwide, while human
fungal pathogens are a major health problem that threatens the lives of millions (6–9).
However, our knowledge of whole reproductive life cycles in the fungal kingdom is only
partial. Sexual reproduction is a key event in the fungal life cycle, where evolution and
selection rely on its creation of novel and beneficial genetic combinations in the organism
(10, 11). The fungal reproductive cycle relies on elaborate genetic regulatory systems as
Citation Brun S, Kuo H-C, Jeffree CE, Thomson
DD, Read N. 2021. Courtship ritual of male and
female nuclei during fertilization in Neurospora
crassa. Microbiol Spectr 9:e00335-21. https://
doi.org/10.1128/Spectrum.00335-21.
Editor Gustavo H. Goldman, Universidade de
Sao Paulo
Copyright © 2021 Brun et al. This is an open-
access article distributed under the terms of
the Creative Commons Attribution 4.0
International license.
Address correspondence to Sylvain Brun,
sylvain.brun@u-paris.fr.
*Present address: Darren D. Thomson,
Department of Biosciences, Medical Research
Council Centre for Medical Mycology at the
University of Exeter, Exeter, United Kingdom.
†Deceased.
Received 3 June 2021
Accepted 25 August 2021
Published 6 October 2021
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 1
RESEARCH ARTICLE
well as complex cytological events (12–14). Understanding reproduction provides means
to mitigate fungal threats and enhance biotechnological processes.
In heterothallic model organisms such as the filamentous fungus Neurospora crassa
and Saccharomyces cerevisiae yeast, sexual reproduction only occurs with opposite
mating-type partners (mat a and mat A in N. crassa) (15). In N. crassa, Dodge identified
the conidium as the male partner and the ascogonium as the female partner (16). The
ascogonium is embedded within a multihyphal female structure called the protoperi-
thecium (17). The ascogonium produces specialized hyphae called trichogynes to facili-
tate fertilization. Trichogynes are chemotropically attracted by a diffusible chemical
signal emitted by conidia (18, 19). Trichogynes eventually fuse to their macro- or micro-
conidial mate, allowing successful fertilization and development of the protoperithe-
cium into a perithecium. Importantly, the nuclei from both parents do not undergo
karyogamy within the trichogyne (20–22).
Trichogynes express G protein-coupled receptors (GPCRs) at their surface which
respond to the pheromone signal emitted by conidia (23–29). In ascomycetes, the
expression of both the pheromone and the GPCRs are under the control of the mat
locus (13). Trichogynes, which are attracted by the pheromone signal bind to, coil
around, and eventually fuse with male conidia, eventually establishing plasmogamy.
Although the migration of male nuclei has never been observed, it is assumed that
once within the female trichogyne, male nuclei (of the opposite sex mating type)
migrate up the trichogyne to the ascogonium, where they proliferate together with
female nuclei, eventually forming the dikaryotic bag (30). Karyogamy and meiosis take
place once two nuclei of each mating type exit the dikaryotic bag to enter the ascoge-
nous cell. Although fertilization and development of perithecia have been imaged in
several ascomycete models, merely by electron micrographs (16, 20, 31–35), the actual
behavior of male and female nuclei in this fundamental process has never been
tracked in living cells.
In the lab, fertilization in N. crassa can be observed a few hours after inoculating the
protoperithecia with a suspension of opposite mating-type conidia, making this fungus
a well-suited experimental model to study fertilization (18–20). Here, using parental N.
crassa strains in which nuclei were visualized with either the synthetic green fluores-
cent protein (H1-sGFP) (36) or the tdimer red fluorescent protein (H1-RFP) fused to the
histone H1 protein, we achieved for the first time live-cell imaging of male and female
nuclei during fertilization. Our observations highlight remarkable differences in the
behavior of nuclei from different parental origins within the same cellular cytoplasm.
Moreover, we imaged the astounding dynamism of male nuclei as they migrate by
repeated stretching and contraction up the trichogyne, passing by immobilized female
nuclei. Finally, our results highlight the intriguing signaling network controlling these
movements of both male and female nuclei throughout fertilization. In particular, our
observations led us to hypothesize that a system preventing the protoperithecium
from being fertilized by several male nuclei in N. crassa may operate similarly to sys-
tems avoiding polyspermy in mammals (37).
RESULTS
Chemotropic growth of trichogynes. To visualize the fertilization process in N. crassa,
we adapted existing experimental settings (18). This involved inoculating 1.5 by 1.5 cm
agar blocks bearing 5- to 7-day-old protoperithecia producing “female”trichogynes with a
freshly prepared (0- to 5-day-old) mixed suspension of “male”macro- and microconidia
from the opposite mating type. After 6 h of inoculation, the agar blocks were inverted onto
a micro-slide chamber containing a droplet of classic Vogel’s medium and imaged on either
an inverted Nikon TE-2000 wide-field or Leica SP8x confocal microscope. Crosses were per-
formed in both directions—$mat A #mat a as well as $mat a #mat A.Weobserved
no differences between the two cross types. Protoperithecia develop on mycelium during
the stationary phase, where growing vegetative hyphae are rare. Within the mycelium net-
work, trichogynes were identified via the following morphological features (Movies S1 and
Brun et al.
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 2
S2;allthemoviesaredownloadablefromhttps://figshare.com/s/e3c33f69ef0131ca6670
[Please do not play them directly online since they do not all play. Download them before
playing them.]) reaching several hundred micrometers (Fig. 1,2,and6);exhibitingtropic
and sinusoidal growth toward conidia (Fig. 1B and C and 2 and Movies S1 and S2) at a
reduced extension rate of 1.1 60.2
m
m/min, compared to vegetative hyphae at the colony
edge (67
m
m/min)(38),andexhibitedcoiledgrowtharoundconidia.Inadditiontotheob-
servation that vegetative hyphae did not exhibit sinusoidal growth (data not shown), tricho-
gynes were only distinguished from vegetative hyphae once they had coiled around a co-
nidium of the opposite mating type. The coiling of the trichogyne around the conidium
was the hallmark of “successful”fertilization (Fig. 1C, 4A, 5A, and 6). Although we only show
here evidence for macroconidia, we also observed microconidia attracting trichogynes
(data not shown) as previously shown (18, 19, 21, 22).
Since conidia are randomly spread on trichogyne-bearing agar blocks, the proximity
of protoperithecia to individual conidia or groups of conidia varied. As a result,
FIG 1 Trichogynes and chemotropic growth in N. crassa. (A) A SEM image of a young (3 days old)
protoperithecium emitting a single trichogyne; scale bar = 10
m
m.(B)Time-lapsesequenceofthetropic
growth of two trichogynes (T) toward one isolated macroconidium (Ma) while ignoring other
macroconidia and microconidia (Mi). The trichogynes are not attracted to the microconidia in this field of
view. Scale bar = 10
m
m. (C) SEM image of a 5- 7-day-old protoperithecium, where one trichogyne has
been attracted and coiled around a conidium. *, Probable trichogyne. Scale bar = 100
m
m.
Live-Cell Imaging of Fertilization in Neurospora crassa
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 3
trichogynes from individual protoperithecia were observed to extend and fuse with
conidia located between a few microns to half a millimeter from their site of origin. We
observed very long trichogynes (.500
m
m) from a protoperithecium fusing with a co-
nidium located close to another protoperithecium. As observed in Fig. 1B, trichogynes
do not necessarily fuse with the first conidium encountered but can instead be
attracted to other conidia further away. Interestingly, an isolated singular macroconi-
dium, in the vicinity of clusters of macroconidia and microconidia, attracted two tricho-
gynes, suggesting heterogeneity in the conidial ability to attract trichogynes (Fig. 1B).
Behavior of female nuclei in the course of fertilization. We imaged H1-sGFP
tagged female nuclei (i) during the tropic growth of trichogynes toward male conidia,
(ii) after the trichogyne had coiled the conidium and received the first mat a H1-RFP
male nucleus, and (iii) when male nuclei migrate up to the trichogyne base. The first
contact between conidia and trichogynes was regularly observed 4 to 6 h after inocula-
tion with the conidial suspension. Entry of male nuclei into trichogynes and protoperi-
thecia occurred 6 to 8 h and 8 to 10 h after inoculation, respectively.
Importantly, female and male H1 histone proteins, fused with either green
(syntheticGFP; sGFP) or red (tdimerRed; RFP) fluorescent proteins, never mixed in the
same nuclei. This enabled us to exclusively follow the GFP-female and RFP-male nuclei
throughout the fertilization process in our imaging experiments. Furthermore, we
never observed karyogamy (fusion of a male and a female nucleus) within trichogynes.
Female nuclei were located and aligned throughout the trichogyne axis (Fig. 2 and
Movie S2). During tropic trichogyne growth, GFP female nuclei displayed an overall an-
terograde movement at rates similar to the extension rate of the trichogyne. From
Fig. 2 and Movie S2, upon trichogyne-conidium contact (time [t] = 36 min), it took
23 min for the trichogyne to complete conidium-coiling and cease growth (t= 59 min).
Next, the first H1-RFP conidial nucleus took 53 min to enter into the trichogyne (t=1 h
FIG 2 Time-lapse imaging of male and female nuclei before and after plasmogamy (Movie S2). Female mat A H1-sGFP was fertilized with mat a H1-RFP
male conidia. (A) merged fluorescence and bright-field confocal images (maximum intensity projected Z-stacks) during the initial stages of fertilization. The
female trichogyne bearing sGFP nuclei tropically grows to one macroconidium containing RFP nuclei (arrowhead). Two RFP male nuclei from the
macroconidium (arrows 1 and 2) enter and migrate up the female trichogyne. (B) GFP fluorescence and bright-field confocal images (maximum intensity
projected Z-stacks) during the later stages of fertilization after RFP male nuclei have left the field of view. Female sGFP nuclei undergo retrograde
movement up the trichogyne from its coiled tip (arrowhead). T, trichogyne. Scale bar = 40
m
m.
Brun et al.
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 4
52 min), followed by a second one 10 min later (t= 2 h 2 min), indicating that plasmog-
amy had occurred. Upon cessation of trichogyne growth at its targeted conidium, the
anterograde movement of female nuclei switched to an oscillatory motion and then
became visually immobilized 4 min before the entry of the first male nucleus (t=1h
48 min). Nucleus speed measurements revealed that female nucleus movements did
not fully stop but were reduced (Fig. 3). For the sake of simplicity, we will refer to this
phase as female nuclear immobilization. These female nuclear dynamics suggest step-
wise signaling events anticipating the entry of the male nucleus. Female nuclei
remained immobilized while male nuclei migrated up the trichogyne. Finally, 56 min
after the first male RFP nucleus entered the trichogyne, all immobilized female GFP
nuclei synchronously began moving backward (retrograde movement) up the tricho-
gyne (t= 2 h 44 min). These female nuclear dynamics were consistent with every fertil-
ization event analyzed (n=.10). We quantified female nucleus mobility before and af-
ter entry of male nuclei into trichogynes for at least one fertilization event (Fig. 3). By
tracking nuclei in Movies S3a (before entry) and S3b (after entry), we measured the
speed of GFP female nuclei within trichogynes after trichogyne growth had ceased at
the conidium and after female nuclei were immobilized. These dynamics were com-
pared to the mobility of female nuclei located in surrounding hyphae, likely vegetative
ones since initial nucleus behavior in these hyphae were different (Fig. 3 and Movies
S3a and b).
Theaveragespeedofhyphalfemalenucleiwassignificantly higher (2.04 62.36
m
m/
min) than that of trichogyne female nuclei (1.03 61.00
m
m/min; Fig. 3; P,10
23
)priorto
male nucleus entry. Strikingly, the speed of both hyphal and trichogyne female nuclei sig-
nificantly decreased after male nucleus entry, to 1.33 61.07
m
m/min (P,10
23
)and
0.82 60.86
m
m/min (P,10
23
), respectively. These data confirmed the immobilization of
female nuclei observed immediately prior to entry of male nuclei into the trichogyne. Note
that the standard deviation (SD) values highlight the variations of individual nuclear move-
ments at each time frame.
FIG 3 Female nuclear speeds before and after male nucleus entry into the trichogyne. Analysis of the
time-lapse Movies S3a and b. (A) Plot; (B) numerical values. The speed of the green-labeled H1-sGFP
female nuclei of the trichogyne and of the vegetative hyphae in the same field of view were
measured at every time frame before (Movie S3a) and after (Movie S3b) entry of a male nucleus into
the trichogyne. S.D, standard deviation; N
nuclei
, number of analyzed nuclei in the field of view; N
speed
,
number of nuclei speeds measured. Results from the ANOVA (on log values) indicate that the effect
of target site is significant (P,0.001), with larger values in hyphae (1.56 61.63, n= 683) than in
trichogyne (0.92 60.94, n= 749); likewise, the effect of time of observation is significant (P,0.001),
with larger values before (1.42 61.73, n= 569) than after (1.10 61.01, n= 863); these two effects
are independent of one another as confirmed by the nonsignificant interaction (P= 0.431). Pairwise t
tests with pooled SD indicate that all pairs of means were different at the 5% level.
Live-Cell Imaging of Fertilization in Neurospora crassa
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 5
The shape of female H1-sGFP fluorescent nuclei in both the analyzed hyphae and in the tri-
chogyne changed from ovoid to more rounded during their immobilized phase, as observed
in Movies S3a and b, and reverted to ovoid at the resumption of movement (end of Movie
S5). During the latter phase (Movie S5), the recovered mobility in female nuclei followed by
their retrograde migration up the trichogyne was exemplified by the behavior of the nucleus,
close to the conidium in Movie S5 (the top one). This exemplar nucleus became pear-shaped
while being pulled in the retrograde direction. These data describe for the first time the
dynamic nature of resident female nuclei within trichogynes during sexual fertilization in
N. crassa. Moreover, the signal that triggers female nuclear immobilization is not restricted
to the trichogyne subjected to plasmogamy, suggesting a possible diffusible signal, which
can pause the nuclear dynamics of this filamentous fungus.
Nuclear morphology and movement of residual conidial male nuclei. Using the
protocol described previously, we aimed to image mat a H1-RFP male nuclei at the onset
of their entry into female mat A H1-sGFP trichogynes. At 4 to 6 h after inoculation with
conidia of the opposite mating type, scarce trichogynes coiling around conidia (fertilization
events) were observed on inoculated agar blocks. In order to image early steps of male nu-
clear migration, image acquisitions were started as soon as coil fertilization hallmarks were
found. This way, we observed that male RFP nuclei were highly mobile, rotating while still
enclosed within the conidium (Fig. 4 and 5; Movies S4 and S5). Eventually, these rotating
movements accelerated until the nucleus (sometimes one of several within the macroconi-
dium) started to repetitively elongate toward the entry of the trichogyne before finally
migrating up the trichogyne. These sequential morphology changes were reproducibly
observed prior to the onset of male nuclear migration.
However, regarding the nucleus imaged in Fig. 4 and the second nucleus remaining
in the conidium (after migration of the first one) in Fig. 5 (Movies S4 and S5), both
FIG 4 Time course of male nuclear mobility within the conidium upon fertilization (Movie S4). (A)
Wide-field fluorescence and bright-field images of H1-sGFP female trichogyne fertilization by an H1-
RFP conidium. (B) Higher magnification of the residual RFP conidial nuclei (box in A) after the first
nucleus entered and migrated up the trichogyne (not shown). The imaged residual nucleus stretched
and moved around within the conidium but failed to migrate up the trichogyne. Time scale = min:
sec. Scale bars = 10
m
m in panel A and 5
m
m in panel B.
Brun et al.
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 6
failed to enter the trichogyne. Both observations of male nuclei failing to engage
migration were representative of other live-cell recordings, where fluorescently imaged
male nuclei did not enter the trichogyne (data not shown). This led us to hypothesize
that the lack of male nuclear migration during live-cell imaging may be due to photo-
toxicity and that real-time frame-rate acquisitions had to be limited prior to migration
observation.
The inch-worm movement of male nuclei. To minimize phototoxicity to the fertil-
ization process, fertilization events were checked infrequently in order to preserve the
integrity of subsequent live-cell imaging of the male nuclear migration. This extra care
allowed observation of several male nuclear migration events into trichogynes. The tri-
chogyne entry process was not straightforward; male nuclei do not directly enter the
trichogyne but, instead, paused frequently prior to entry. It was frequently observed
that individual male nuclei stretched and retracted in and out of the trichogyne several
times without leaving the conidium (data not shown). This “inch-worm”nuclear move-
ment is illustrated in Fig. 5 and Movie S5, where the RFP male nucleus moved in an
FIG 5 Time course of male nucleus movement across the trichogyne (Movie S5). (A) Merged confocal images (maximum-intensity projected Z-stacks) of the
fertilization of a mat A H1-sGFP female trichogyne by a mat a H1-RFP conidium. The RFP nucleus enters and migrates up the female trichogyne
successively, stretching and contracting in an inch-worm-like manner. The remarkable stretching (40
m
m) of the male nucleus crossing the first septa
(arrowheads) is indicated. Scale bar = 5
m
m. (B) Velocity analysis of the exemplar stretching RFP nucleus at the front (gray) and rear points (black).
Live-Cell Imaging of Fertilization in Neurospora crassa
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 7
“inch-worm”manner, repetitively stretching and contracting while migrating up the
trichogyne. Stretching was so dramatic that male nuclei seemed sometimes split into
two (Fig. 5; from 3 min 18 s to 4 min 3 s; Movie S5). The way male nuclei stretched sug-
gested that the front is submitted to pulling mechanical strain during migration (see
Discussion). Nuclear elongation was especially striking when the male nucleus passed
through septa. In Fig. 5, the male nucleus passed through the first septum (Fig. 5;
t= 2 min 10, s to t= 4 min 3 s) and, remarkably, stretched to a maximum length of
40
m
m(t= 3 min 27 s; Fig. 5), where its apical end was pulled while its rear end was
“stuck”at the septal pore. To understand this process more, we measured the speed of
each nuclear apex (front and rear) in the movie. The maximum speed measured for the
front side was 85
m
m/min, while the maximum speed of the rear side reached
130
m
m/min (Fig. 5B). These data highlight the remarkable spatiotemporal dynamics
and nature of migrating fungal nuclei during fertilization in N. crassa.
Fertilization of the protoperithecium by male nuclei. After trichogyne entry, the
conidial nuclei travel to the protoperithecium, where the next steps of sexual repro-
duction, such as karyogamy, meiosis, and ascosporogenesis, take place. In N. crassa,
macroconidia are typically multinucleate. We reproducibly observed that fertilization
by macroconidia led to the discharge of several nuclei from a single conidium into a tri-
chogyne (i.e., three nuclei per conidium in Fig. 6, 2 nuclei in Fig. 2, and 2 nuclei in
Movies S7 and S8). Furthermore, we observed that trichogynes can branch and that
those branches are capable of binding, coiling, and fusing with at least two conidia (A
and B; Fig. 6 and Movie S6). In the trichogyne of Fig. 6, male nuclei were tracked from
the conidia to the protoperithecium over a distance of up to 574
m
m (Fig. 6; conidium
A). Three male RFP nuclei entered the trichogyne from conidium A in 35 min (Fig. 6, la-
beled 1, 2, and 3; from t= 46 min to t= 1 h 21 min), followed by three additional nuclei
from conidium B, which were released in only 6 min (Fig. 6, labeled 4, 5, and 6; from
t= 1 h 25 min to t= 1 h 31 min). By carefully tracking the different male nuclei within
the trichogyne, we determined that the nucleus entering the protoperithecium was
nucleus 4 from conidium B (Fig. 6; t= 1 h 53 min). Strikingly, although the remaining
male nuclei continued their migration up the trichogyne, all of them stalled and accu-
mulated upstream of the now-fertilized perithecium (Fig. 6; t= 4 h 24 min; Movie S6).
It was also the case in Movie S7, where two conidial male nuclei migrated up the tri-
chogyne and only one entered the protoperithecium. Accordingly, immobile RFP male
nuclei in trichogynes were frequently observed late in the experiment (between 10
and 14 h), where presumably, nuclei were somehow blocked from migrating after pro-
toperithecium fertilization (data not shown). Note that the two full fertilization events
imaged (Fig. 6, Movies S6 and S7) were the only recordings where we tracked multiple
male nuclei migrating into a trichogyne from plasmogamy to entry into a protoperithe-
cium. These live-cell recordings led us to hypothesize that entry of a first male nucleus
into a protoperithecium inhibits entry of the following nuclei.
A second important feature provoked by entry of male nuclei into prothoperithecia
was the quantified increase of volume of the latter (Fig. 6; from t= 1 h 53 min; Fig. 7).
Finally, we detected a second RFP focal signal in the core of the protoperithecium
twice (Fig. 6; t= 2 h 25 min to 4 h 27 min; Movie S7; 4 h 22 min 52 sec). Careful analysis
of the images did not indicate any entry of any of the remaining nuclei. Thus, the
appearance of this second RFP focal signal 32 min after initial protoperithecium entry
in Fig. 6 (Movie S6) may indicate that this male nucleus 4 has divided. However, we
cannot exclude that the appearance of a second focus within the protoperithecium
FIG 6 Legend (Continued)
first row shows merged GFP female nuclei and RFP male nuclei with a bright-field representation at
t= 0 min. Subsequent time-lapse RFP images contain a dotted trace of the trichogyne and
protoperithecium. Two conidia (A and B) attracted and fused with a branched trichogyne emitted by
the protoperithecium at the bottom-right corner. Three nuclei (1, 2, and 3) from conidium A and three
nuclei (4, 5, and 6) from conidium B successively enter the trichogyne. Nucleus number 4 enters the
protoperithecium core and eventually divides (4a and 4b). Scale bar = 40
m
m; time scale = h:min:sec.
Live-Cell Imaging of Fertilization in Neurospora crassa
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may be due to the stretching of the nucleus. These data suggest that entry of male
nuclei into protoperithecia may trigger at least two signals, a first one in order to avoid
polyspermy by blocking multiple fertilization of the protoperithecium and a second
signal committing the protoperithecium into a growing and developing perithecium.
The orienteering race of male nuclei up the trichogyne. Another astonishing
behavior of conidial male nuclei is their capacity to rapidly switch between antero-
grade and retrograde movement during trichogyne migration (Movies S8 and S9).
Moreover, trichogynes with branches had male nuclei switching direction at T-junc-
tions, illustrated in Movie S8 (the RFP male nuclei are colored in yellow in this movie),
where the first male nucleus initially migrated right but then reversed and migrated
left toward its intended pathway to the protoperithecium. This direction switch of
male nuclei at T-junctions has been observed several times in independent experi-
ments (data not shown). These observations led us to conclude that male nuclei are
endowed with the capacity to change direction in the course of migration and to per-
haps follow a yet unknown signal toward the protoperithecium.
An interesting feature of trichogynes is their ability to fuse with several conidia and,
as a consequence, to allow the discharge of nuclei belonging to different conidia
(Fig. 6 and Movie S6). It is interesting to note that nuclei from conidium A stopped
migration, especially nucleus 1, remaining immobile for 23 min (from t= 1 h 25 min
58stot= 1h 52 min 59 s), while the nuclei from conidium B were released into the tri-
cogyne. We measured the average speed of nuclei 1 and 2 before the pause and com-
pared it to their average speed all along the whole trichogyne. Before the pause and
across the first 420
m
m of migration, the speeds of nuclei 1 and 2 were 10.9 and
12.3
m
m/min, respectively, compared to 6.9 and 8.0
m
m/min, respectively, for the
entire migration of both nuclei. This indicated that the speed of male nuclei is not con-
stant during their migration all along the trichogyne and that migration of some male
nuclei may interfere with the migration of other nuclei within the same trichogyne.
DISCUSSION
Sexual reproduction in fungi and fruiting body development have attracted the in-
terest of researchers for centuries (39). Imaging these structures under non-live-cell
imaging conditions, such as electronic microscopy, has highlighted the extraordinary
complexity of fruiting bodies (31–34). However, the lack of live-cell biology has been
an obstacle to fully understanding sexual reproduction from the initial plasmogamy of
sexual cells to the final production of sexual spores. In previous Neurospora studies, fol-
lowing the work of B. O. Dodge, who identified the sexual reproductive organs in the
FIG 7 Volume increase of protoperithecium after its fertilization by entry of one male nucleus.
Analysis of segmented images of H1-sGFP fertilization by H1-RFP conidial suspension in Movie S6
(Fig. 6). The green signal of the H1-sGFP nuclei composing the protoperithecium was segmented in
order to evaluate the volume of the perithecium in the course of fertilization. The time of entry of
the first H1-RFP male nucleus into the trichogyne (plasmogamy) and the time of entry of the nucleus
number 4 into the protoperithecium (protoperithecium fertilization) are indicated. Dots, individual
data points; black line, moving average.
Brun et al.
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 10
Neurospora genus, Backus in 1939 observed the fertilization of trichogynes by conidia
and described it through incredibly precise drawings (15, 20). In particular, Backus
observed that trichogynes could be branched and were able to bind to and coil around
(presumably fuse with) several conidia of the opposite mating type (20). Almost a cen-
tury after these pioneering studies, using modern live-cell imaging technologies, we
are able to image live conidial fertilization. Through live-cell imaging, we visualized the
stages of N. crassa ontogeny for the very first time, encompassing the migration of
male nuclei along the entire trichogyne length from their parental conidia to the asco-
gonium at the core of protoperithecia. Our experimental system contained male and
female partners differentially bearing H1-sGFP or H1-RFP tagged nuclei. Importantly,
we never observed nuclei sharing red and green fluorescent protein, implying that
both these reporter proteins (H1-sGFP and H1-RFP) did not mix once nuclei bearing
each of these transgenes were together within the same trichogyne cytoplasm. This
property makes our experimental model well suited to describe and demonstrate the
striking opposite behaviors of male and female nuclei within the trichogyne and con-
firmed that karyogamy does not take place right after plasmogamy within the tricho-
gyne in this fungus (21, 22, 30).
During trichogyne growth, resident female nuclei undergo anterograde movement
at a comparable speed to the trichogyne. Whether this movement is supported by
cytoplasmic flow as in vegetative hyphae and whether female nuclei divide during tri-
chogyne growth remain to be addressed (40). Upon cessation of trichogyne growth at
the male conidium, female nuclei occupy the entire length of the trichogyne and are
clustered regularly, especially at the trichogyne tip. After contact between the tricho-
gyne and the conidium, the female nuclei only show oscillatory movements. In a strik-
ingly concerted manner, movements of female nuclei stop right before entry of the
first male nucleus into the trichogyne and eventually resume, showing female nuclei
migrating backward toward the protoperithecium. Identifying what controls female
nuclear movements during fertilization will be a great challenge. The first clues are
given by the observation of female nuclear shape during the different steps described
above. Indeed, female nuclear shape changes from oval or pear-shape before entry of
male nuclei to round during the immobilization phase and then oval or pear-shaped
when they move again. These changes may be reminiscent of modifications in the
interaction between these nuclei and the cytoskeleton. In filamentous fungi, the micro-
tubule (MT) cytoskeleton, as well as its associated molecular motors, i.e., the kinesins
and the dynein-dynactin complex, have been involved in finely tuned movements
such as shape changes, oscillatory movements, nuclear division, and nuclear distribu-
tion in hyphae. In particular, both changes of shape and oscillatory movements are
affected when cells are treated with MT-depolymerizing drugs as well as in the ropy
mutants altered for the dynein-dynactin complex (36, 41–44). Moreover, nuclei in the
ropy mutants and in the kin-1 mutant (altered for the major molecular motor Kinesin-1)
have a rounded shape, in contrast to the oval or pear shape of nuclei in wild-type cells.
Therefore, it is likely that the oscillatory movement of female nuclei within the tricho-
gyne might involve the MT cytoskeleton and the kinesin/dynein molecular machinery.
We speculate that the change of shape of female nuclei together with their immobili-
zation may be reminiscent of momentary loss of interaction of these nuclei with the
MT cytoskeleton machinery during the time when male nuclei migrate. In addition, we
have observed that the immobilization of female nuclei may not be trichogyne-auton-
omous, since it was detected in surrounding hyphae. Altogether, these data led us to
hypothesize that a diffusible signal that triggers immobilization of female nuclei is pro-
duced at the onset of male nuclear entry into the trichogyne.
In contrast to the resident female nuclei, male nuclei have to migrate up the tricho-
gyne in order to reach the ascogonium in the core of the protoperithecium. Here, this
assumption has been proven correct via the tracking of male nuclei all along their
migration across the trichogyne until the core of the protoperithecium, presumably
the ascogonium. We observed that several male nuclei could enter a single trichogyne,
Live-Cell Imaging of Fertilization in Neurospora crassa
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 11
raising the question of the poly-fertilization of protoperithecia. The possibility for more
than one male nucleus to fertilize a single protoperithecium has genetically been
addressed in N. crassa (45–47). Sansome was the first to demonstrate that ascospores
from a single perithecium could originate from more than one pair of parental nuclei.
In 1960, Weijer (47) showed that 23% of the perithecia contained rosettes initiated by
more than one male nucleus when a heterokaryotic male parent was used. However,
Nakamura et al. (45) found that only 2% (60/2,770) of perithecia display mosaic rosettes
accounting for fertilization by two or more different male nuclei when a mix of geneti-
cally different conidia is used. This shows that when using this more “natural”method
(very similar to our fertilization conditions), fertilization of a single protoperithecium by
more than one male nucleus is a rare event.
Here, we show for the first time that several male nuclei can indeed enter a single
trichogyne and that they can originate from different conidia. Furthermore, we often
observed branched trichogynes, suggesting that fertilization of a protoperithecium by
more than one male parent is feasible, albeit likely rare (45). However, in the two cases
for which we tracked several male nuclei migrating as far as the protoperithecium, we
observed that only one male nucleus penetrated the core of the protoperithecium,
while the following nuclei within the same trichogyne remained outside. Although fur-
ther experiments will be required to confirm these observations, we speculate that
entry of the first male nucleus into the protoperithecium may inhibit entry of the fol-
lowing ones. If such a blockage exists, it may be leaky, thus facilitating poly-fertilization
of protoperithecia in rare cases (45). This blockage mechanism in the fungus N. crassa
may be reminiscent of the mechanism in mammals which avoids “polyspermy.”
Although we ignore how male nuclei find the way to the protoperithecium in a
branched trichogyne, we observed male nuclei first making a hesitant turn at T-junc-
tions and then reversing to take the other branch, likely the correct path to the proto-
perithecium. This highlights that male nuclei can rapidly change direction in a tricho-
gyne (moving back and forth in a retrograde versus an anterograde movement) and
that the migration path is not unique, nor is its signaling precisely defined. We specu-
late that this signaling may regulate the tethering of male nuclei to molecular motors
but, also, the choice of the appropriate molecular motor to move in the anterograde or
the retrograde direction.
The most striking features observed in our attempts to characterize the behavior of
nuclei in the course of fertilization are indubitably the ones related to male nuclei. Male
nuclei undergo a series of morphological changes within the conidium at the onset of
migration. Since we could not observe when cell fusion (plasmogamy) between tricho-
gynes and conidia occurs, how these morphological changes in the nuclei synchronize
with plasmogamy remains to be addressed. Next, male nuclei undergo initial stretching
upon leaving the conidium. Male nuclei then migrate up the trichogyne, repetitively
stretching and contracting in an inch-worm-like manner. Generally, the migrating nuclei
harbor a compact shape before encountering septa and elongate dramatically while pass-
ing through. This stretching can be as extreme as 40
m
m in length, around 20 times the
size of standard nuclei (2
m
m), shedding light on two important features of migrating
male nuclei—the extraordinary plasticity of the nuclei and the strong pulling force applied
to the nucleus. This plasticity is particularly striking when the rear of the nuclei remains
blocked at septa while, in the meantime, the front is repetitively pulled, leading to repeti-
tive stretching (Movie S5). In the end, like a rubber band stretched and released at one
side, the rear of the nucleus eventually joins the front in a movement exerting one of the
highest speeds ever measured for nuclei in cells (130
m
m/min). We propose that the force
moving nuclei in the trichogyne is likely a pulling force applied to the front of the nucleus
and that the rear follows by a spring effect. Since male nuclei can rapidly change direction
and move back and forth, this implies that the leading “front”of nuclei can become the
“rear”and vice-versa almost instantly.
Regarding the male nuclear velocity, its versatility, and the fact that during this im-
pressive mobility, female nuclei remain immobilized, we exclude the cytoplasmic flow
Brun et al.
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 12
to be responsible for male nuclear movements. Hence, we suppose that the move-
ments of male nuclei are undertaken by molecular motors and the cytoskeleton (41).
Further studies are required to determine the role of the microtubules and actin, as
well as the role of their cognate molecular motors, i.e., myosin versus dynein/dynactin
and kinesin, respectively, in the movement of male (but also of female) nuclei, within
the trichogyne during sexual reproduction.
Nuclear movements and deformation have been studied in many organisms (plants,
animals, and fungi) and in many contexts (during development, sexual reproduction,
hematopoiesis, metastases, migration, etc.). No such deformation or speeds have been
reported in plants or animals, where one the highest speeds measured was 16
m
m/min
for the male pronucleus during fertilization in Xenopus (reviewed in reference 48).
Interestingly, high nuclear speeds have been reported in N. crassa during hyphal
growth (40). At the hyphal apex, nuclei migrate at a speed of 8 to 9
m
m/min, principally
propelled by bulk flow. These nuclei also show very rapid and minute anterograde/ret-
rograde movements that can reach 74.4
m
m/min for anterograde movements in the
wild-type and even 222
m
m/min for retrograde movements in a ro-1 dynein mutant
(40). These data show that N. crassa is endowed with a molecular machinery enabling
very rapid nuclear movements, and it is likely that a similar machinery is involved in
rapid male nuclear movements during their migration in trichogynes.
The difference in behavior of male and female nuclei inside the same fungal cell
cytoplasm raises the question of the identity of both types of nuclei. Since heterothallic
fungi can only reproduce by sexual reproduction with individuals of the opposite mat-
ing type, it is assumed that the mating type locus is the primary determinant of the
male and the female nuclear identity (13). In particular, mat loci regulate the expres-
sion of the pheromones and their cognate GPCRs, which are both involved in the che-
motropic interplay between conidia and trichogynes. Basically, binding of the phero-
mone produced by a conidium to its cognate receptor at the trichogyne membrane
activates the GPCR signaling cascade responsible for the chemotropic growth, the coil-
ing, and ultimately, the fusion with conidia of the opposite mating type only (25, 26,
28, 29). It will be of great interest to test whether the mat locus and the pheromone/
GPCR pathway are involved in the control of male and female nuclear movements dur-
ing sexual reproduction in N. crassa.
Through modern live-cell imaging, our work humbly aimed to answer questions
about nuclear fate in the course of fertilization as old as the discovery of sex in fungi
(39). However, our study broadens the perspectives beyond the simple study of fertil-
ization by unravelling unexpected behavior of both male and female nuclei. This live-
cell imaging approach to studying fertilization in the model ascomycete N. crassa raises
new questions. What are the regulation pathways that control immobilization of
female nuclei? What is the machinery that pulls male nuclei during migration? What
signal guides male nuclei during migration? Is there a polyspermy avoidance, and how
is it set up? How does male nuclear entry into protoperithecia trigger perithecial devel-
opment? What defines nuclear identity? On top of this, the dramatic deformation of
male nuclei during migration within the trichogyne and the amenability of model fungi
such as N. crassa for live-cell biology and molecular genetics studies make fertilization
in N. crassa a new system to use to study the effect of physical forces and the deforma-
tions they create on nuclei. Understanding how physical forces and constraints applied
to nuclei modify the whole nucleus organization and trigger gene expression switches
is an emerging area of research (49, 50). The number of human pathologies primarily
due to failure in mechanotransduction of signals is constantly increasing (51), and
recent data show that extreme deformation of nuclei in metastatic cells during migra-
tion has a priming effect in aggressiveness of these malignant cells (52). Therefore, the
study of the plasticity of the male nucleus and of its chromatin structure during sexual
reproduction in N. crassa offers a unique opportunity to unravel how extreme deforma-
tions of nuclei can modulate genome structure and gene expression in eukaryotes.
Live-Cell Imaging of Fertilization in Neurospora crassa
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 13
MATERIALS AND METHODS
Strains, culture conditions, and production of conidia. The Neurospora crassa strains used in this
the study were wild-type strains 74-OR23-1V A (FGSC no. 2489) and ORS-SL6 a (FGSC no. 4200), N2282
mat A his-3
1
::Pccg-1-hH1
1
-sGFP (H1-sGFP in the article), N2283 mat a his-3
1
::Pccg-1-hH1
1
-sGFP, and mat
a his-3
1
::Pccg-1-hH1
1
-tdimerRed (H1-RFP in the manuscript). Construction of the H1-RFP strain (no. 263 in
the Nick Read collection) was conducted as follows: a plasmid expressing an N-terminal tdimerRed fluo-
rescent protein (RFP) fused to histone H1 was constructed by inserting the N. crassa hH1 gene, amplified
from pMF280 (36) with primers hH1SpeF (59-GCCACTAGTATGCCTCCCAAGAAGACCGAG-39) and
hH1XbaR (59-GCCTCTAGATTGCCTTCTCGGCAGCG-39) into pMF331 (53) digested with SpeI and XbaI. The
resulting plasmid, pMF360, was transformed into N623 (mat A his-3 [36) as previously described (54).
Positive transformants were selected on minimal medium and screened for expression of tdimerRed-H1
(H1-RFP in the article) under a dissecting fluorescence microscope as previously described (53). A trans-
formant (NMF138) was crossed with N3011 (mata his-2;mus-51::bar1) to generate homokaryotic prog-
eny. One mat a progeny was called strain 263 in the Nick Read collection.
Strains were maintained and grown on solid classic Vogel’s minimal medium with 2% sucrose at
27°C. Conidia were harvested from 4- to 5-day-old cultures grown at 25°C on Vogel’s medium with con-
stant light, filtered with Miracloth (Merck Millipore), washed in TS (0.05% Tween 80/9% NaCl/sterilized
water), and stored in TS at 4°C for up to 5 days.
Fertilization procedure. Petri plates containing 2% agar (tap water) solid medium were prepared
and inoculated with the female strain (mat a or A) and incubated for 5 to 7 days until protoperithecia
developed. A 2- to 3-cm
2
agar plug harboring protoperithecia was cut and transferred to an empty petri
plate. This sample was inoculated with a 1 to 2.10
6
conidia Ml
21
suspension of the “male”partner of
the opposite mating type. Conidial suspensions were prepared the same day as the inoculation or kept
at 4°C in TS before use (up to 5 days). After 4 to 6 h of incubation at 25°C in a moisture chamber, the
sample was mounted inverted onto a droplet of Vogel’s medium in a 2-well
m
-Slide ibidiTreat chamber
and inspected for fertilization. Alternatively, after the initial four hours of incubation at 25°C, the sample
was kept at 4°C overnight and mounted the next day in the microscopy chamber. In both cases, migrat-
ing male nuclei were observed after 2 to 5 h of further incubation at room temperature.
Low-temperature scanning electron microscopy (LTSEM). For Fig. 1A and C, low sucrose agar
(LSA) plates overlaid with sterile cellophane (525-gauge uncoated Rayophane; A.A. Packaging, Preston,
UK) were used for sample preparation. Fig. 1A shows 3-day-old culture of the wild-type mat A N. crassa
strain (74A). Figure 1C shows the 7-day-old wild-type (74a) N. crassa strain inoculated with wild-type mat
A male conidia (74A) (14 h of incubation after inoculation). Cellophane rectangles carrying the specimen
were cut out and attached to the surface of a cryospecimen carrier (Gatan, Oxford, UK) with a thin layer
of Tissue-Tek OCT compound (Sakura Finetek, Torrance, CA) as an adhesive and cryofixed by plunging
into subcooled liquid nitrogen. The specimen carrier was transferred under low vacuum to the cold
stage of a Gatan Alto 2500 cryo-preparation system at 2180°C and then to the cold stage of an S4700II
field emission scanning electron microscope (Hitachi, Wokingham, UK), where it was warmed to 280°C
under continuous visual observation until any surface ice contamination was removed by sublimation.
The specimen was then recooled to below 2120°C before being returned to the specimen stage of the
Gatan Alto 2500 cryo-preparation system at 2180°C, where it was coated with about 10 nm of 60:40
gold-palladium alloy (Testbourne Ltd., Basingstoke, UK) in an argon gas atmosphere. The coated speci-
men was returned to the cold stage of the SEM and examined at a temperature of ,–160°C, a beam
accelerating voltage of 2 kV, and a beam current of 10
m
A, at working distances of 12 to 15 mm. Digital
images were captured at a resolution of 2,560 by 1,920 pixels by 8 bits using the signal from the lower
secondary electron detector and were saved in TIFF format.
Live-cell imaging. Confocal imaging. Fertilization events were imaged in inverted agar samples on
a Leica SP8x confocal microscope equipped with argon and supercontinuum white light laser (WLL) exci-
tation sources. The excitation wavelengths were 488 nm (sGFP; argon laser) and 555 nm (RFP;WLL); the
emission was collected on HyD detectors for sGFP (standard, 496 to 550 nm) and tdimerRed (standard,
600 to 721 nm), while a transmitted light photon multiplier tube (PMT) was used for bright-field collec-
tion. The acquisition settings were as follows. For Fig. 2, 6 and 7 and Movies S1, S2, S6, and S7, objective,
HCX PL APO CS 10/0.40 dry; 8 stack intervals of 4.3
m
m; stacks aquired every 2 mins 27.4 secs. For
Fig. 4 and 5 and Movies S3a and b, S4, S5, S8, and S9, objective, HCX PL APO UVIS CS2 63/1.20 water.
For Movie S3a, 31 stack intervals of 0.7
m
m; stacks aquired every 1 min. For Movie S3b, 18 stack intervals
of 1
m
m; stacks aquired every 9.9 secs. For Fig. 5 and Movie S5, 16 stack intervals of 0.7
m
m; stacks
aquired every 4.5 secs. For Movie S9, 28 stacks intervals of 1
m
m; stacks aquired every 5.4 secs. For
female nuclear speed measurements, nuclei were segmented into spots with IMARIS software (Oxford
Instruments, UK), and the track speeds for each spot were extracted.
Except for Fig. 1 and 4 and Movie S4, all the pictures and movies were generated with IMARIS as Z-
projections with maximum intensity. Fig. 2, 4, 5, and 6 were done using the function “montage”of FIJI
software (https://imagej.net/Fiji) (55). For Fig. 4 and Movie S4, wide-field imaging was performed on a
temperature-controlled motorized Nikon Te2000 microscope equipped with a Nikon PlanApo VC water
immersion 60/1.2 NA lens objective. Fluorescence images were captured on a Hamamatsu Orca-ER
charge-coupled device (CCD) camera (Hamatsu Photonics UK Ltd., UK) under pE-1 LED illumination
(CoolLED, UK) exciting at 470 and 550 nm for sGFP and tdimerRed, respectively, using a Semrock GFP/
DsRes-2x-A-NTE filter cube (Chroma) and MetaMorph software (Molecular Devices). Four-dimensional
(4D) time-lapse stacks were acquired with 3 slices at a stack interval of 1
m
m with a frame rate of
29.6 sec
21
Brun et al.
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 14
Statistical analysis. A two-way analysis of variance (ANOVA) with fixed effects for target site
(hyphae versus trichogyne), time of observation (before versus after nuclear male entry), and their inter-
action was carried out on all available data. Multiple observations from identical loci were pooled using
the arithmetic mean, and data were log-transformed to mitigate the right tailed distribution and to sta-
bilize the variance. Post hoc tests were corrected for multiple comparison using the Holm stepwise pro-
cedure to ensure an adequate family-wise error rate. A type I error rate of 0.05 was considered for all sta-
tistical analyses.
ACKNOWLEDGMENTS
We dedicate this article to the memory of our beloved dear friend Nick Read.
All imaging and analysis were performed at the Institute of Cell Biology of the
University of Edinburgh and at the Phenotyping Centre at Manchester (PCAM) of the
University of Manchester.
S.B. is an assistant professor employed by the Université de Paris and has received a
grant from the Université de Paris for his sabbatical at the MFIG, where he performed
the experiments. This study was supported by IdEx Université de Paris (ANR-18-IDEX-
0001); H.-C.K. was a Ph.D. student and was funded by Overseas Research Studentship.
We thank Kathryn Lord for the permission to use the picture of the young
protoperithecium in Fig. 1A, Kathryn Lord and Alexander Lichius for their help in finding
the origin of the H1-RFP strain used in this paper, Michael Freitag for providing us with
the details of the H1-RFP construction, Christophe Lalanne for the statistical analyses,
and Vincent Contremoulins from the ImagoSeine microscopy facility for his help
generating Movies S5 and S9; the ImagoSeine facility is a member of the France
BioImaging infrastructure supported by the French National Research Agency (ANR-10-
INSB-04, “Investmentsfit the future”).
S.B.: experimentation, data analysis, and manuscript writing; H.-C.K.: experimentation;
C.E.J.: experimentation; D.D.T.: manuscript writing; N.R.: project leader and manuscript
writing.
We declare no conflict of interest.
REFERENCES
1. Dighton J, Tugay T, Zhdanova N. 2008. Fungi and ionizing radiation from
radionuclides. FEMS Microbiol Lett 281:109–120. https://doi.org/10.1111/j
.1574-6968.2008.01076.x.
2. Onofri S, de Vera J-P, Zucconi L, Selbmann L, Scalzi G, Venkateswaran KJ,
Rabbow E, de la Torre R, Horneck G. 2015. Survival of Antarctic cryptoendo-
lithic fungi in simulated Martian conditions on board the International Space
Station. Astrobiology 15:1052–1059. https://doi.org/10.1089/ast.2015.1324.
3. Paulussen C, Hallsworth JE, Álvarez-Pérez S, Nierman WC, Hamill PG, Blain
D, Rediers H, Lievens B. 2017. Ecology of aspergillosis: insights into the
pathogenic potency of Aspergillus fumigatus and some other Aspergillus
species. Microb Biotechnol 10:296–322. https://doi.org/10.1111/1751
-7915.12367.
4. Cairns TC, Nai C, Meyer V. 2018. How a fungus shapes biotechnology: 100
years of Aspergillus niger research. Fungal Biol Biotechnol 5:13. https://
doi.org/10.1186/s40694-018-0054-5.
5. Chambergo FS, Valencia EY. 2016. Fungal biodiversity to biotechnology.
Appl Microbiol Biotechnol 100:2567–2577. https://doi.org/10.1007/s00253
-016-7305-2.
6. Bongomin F, Gago S, Oladele RO, Denning DW. 2017. Global and multi-
national prevalence of fungal diseases: estimate precision. J Fungi 3:57.
https://doi.org/10.3390/jof3040057.
7. Brown GD, Denning DW, Gow NAR, Levitz SM, Netea MG, White TC. 2012.
Hidden killers: human fungal infections. Sci Transl Med 4:165rv13. https://
doi.org/10.1126/scitranslmed.3004404.
8. Fisher MC, Henk DA, Briggs CJ, Brownstein JS, Madoff LC, McCraw SL, Gurr
SJ. 2012. Emerging fungal threats to animal, plant and ecosystem health.
Nature 484:186–194. https://doi.org/10.1038/nature10947.
9. Köhler JR, Hube B, Puccia R, Casadevall A, Perfect JR. 2017. Fungi that infect
humans. Microbiol Spectr 5:5.3.08. https://doi.org/10.1128/microbiolspec
.FUNK-0014-2016.
10. Heitman J. 2010. Evolution of eukaryotic microbial pathogens via covert
sexual reproduction. Cell Host Microbe 8:86–99. https://doi.org/10.1016/j
.chom.2010.06.011.
11. Heitman J, Sun S, James TY. 2013. Evolution of fungal sexual reproduc-
tion. Mycologia 105:1–27. https://doi.org/10.3852/12-253.
12. Bennett RJ, Turgeon BG. 2016. Fungal sex: the Ascomycota. Microbiol
Spectr 4:4.5.20. https://doi.org/10.1128/microbiolspec.FUNK-0005-2016.
13. Debuchy R, Berteaux-Lecellier V, Silar P. 2010. Mating systems and sexual
morphogenesis in Ascomycetes, p 501–535. In Borkovitch K, Ebbole D
(ed), Cellular and molecular biology of filamentous fungi. ASM Press,
Washington DC.
14. Ni M, Feretzaki M, Sun S, Wang X, Heitman J. 2011. Sex in fungi. Annu Rev
Genet 45:405–430. https://doi.org/10.1146/annurev-genet-110410-132536.
15. Dodge BO. 1927. Nuclear phenomena associated with heterothallism and
homothallism in the ascomycete Neurospora. J Agric Res 35:289–305.
16. Dodge BO. 1935. The mechanics of sexual reproduction in Neurospora.
Mycologia 27:418–438. https://doi.org/10.2307/3754168.
17. Lichius A, Lord KM, Jeffree CE, Oborny R, Boonyarungsrit P, Read ND.
2012. Importance of MAP kinases during protoperithecial morphogenesis
in Neurospora crassa. PLoS One 7:e42565. https://doi.org/10.1371/journal
.pone.0042565.
18. Bistis GN. 1981. Chemotropic Interactions between trichogynes and coni-
dia of opposite mating-type in Neurospora crassa. Mycologia 73:959–975.
https://doi.org/10.2307/3759806.
19. Bistis GN. 1983. Evidence for diffusible, mating-type-specific trichogyne
attractants in Neurospora crassa. Exp Mycol 7:292–295. https://doi.org/10
.1016/0147-5975(83)90051-8.
20. Backus MP. 1939. The mechanics of conidial fertilization in Neurospora
sitophila. Bull Torrey Bot Club 66:63–76. https://doi.org/10.2307/2480991.
21. Davis RH. 2000. Neurospora: contributions of a model organism. Oxford
University Press, New York, NY.
22. Nelson MA, Metzenberg RL. 1992. Sexual development genes of Neurospora
crassa. Genetics 132:149–162. https://doi.org/10.1093/genetics/132.1.149.
23. Bobrowicz P, Pawlak R, Correa A, Bell-Pedersen D, Ebbole DJ. 2002. The
Neurospora crassa pheromone precursor genes are regulated by the mat-
ing type locus and the circadian clock. Mol Microbiol 45:795–804. https://
doi.org/10.1046/j.1365-2958.2002.03052.x.
Live-Cell Imaging of Fertilization in Neurospora crassa
Volume 9 Issue 2 e00335-21 MicrobiolSpectrum.asm.org 15
24. Jones SK, Bennett RJ. 2011. Fungal mating pheromones: choreographing
the dating game. Fungal Genet Biol 48:668–676. https://doi.org/10.1016/j
.fgb.2011.04.001.
25. Kim H, Borkovich KA. 2004. A pheromone receptor gene, pre-1, is essen-
tial for mating type-specific directional growth and fusion of trichogynes
and female fertility in Neurospora crassa. Mol Microbiol 52:1781–1798.
https://doi.org/10.1111/j.1365-2958.2004.04096.x.
26. Kim H, Borkovich KA. 2006. Pheromones are essential for male fertility
and sufficient to direct chemotropic polarized growth of trichogynes dur-
ing mating in Neurospora crassa. Eukaryot Cell 5:544–554. https://doi
.org/10.1128/EC.5.3.544-554.2006.
27. Kim H, Metzenberg RL, Nelson MA. 2002. Multiple functions of mfa-1, a
putative pheromone precursor gene of Neurospora crassa. Eukaryot Cell
1:987–999. https://doi.org/10.1128/EC.1.6.987-999.2002.
28. Kim H, Wright SJ, Park G, Ouyang S, Krystofova S, Borkovich KA. 2012.
Roles for receptors, pheromones, G proteins, and mating type genes dur-
ing sexual reproduction in Neurospora crassa. Genetics 190:1389–1404.
https://doi.org/10.1534/genetics.111.136358.
29. Krystofova S, Borkovich KA. 2006. The predicted G-protein-coupled recep-
tor GPR-1 is required for female sexual development in the multicellular
fungus Neurospora crassa. Eukaryot Cell 5:1503–1516. https://doi.org/10
.1128/EC.00124-06.
30. Raju NB. 1992. Genetic control of the sexual cycle in Neurospora. Mycol
Res 96:241–262. https://doi.org/10.1016/S0953-7562(09)80934-9.
31. Harris JL, Howe HB, Roth IL. 1975. Scanning electron microscopy of surface
and internal features of developing perithecia of Neurospora crassa. J Bacter-
iol 122:1239–1246. https://doi.org/10.1128/jb.122.3.1239-1246.1975.
32. Lord KM, Read ND. 2011. Perithecium morphogenesis in Sordaria macro-
spora. Fungal Genet Biol 48:388–399. https://doi.org/10.1016/j.fgb.2010
.11.009.
33. Mai SH. 1976. Morphological studies in Podospora anserina. Am J Bot 63:
821–825. https://doi.org/10.2307/2442040.
34. Read ND. 1983. A scanning electron microscopic study of the external fea-
tures of perithecium development in Sordaria humana. Can J Bot 61:
3217–3229. https://doi.org/10.1139/b83-359.
35. Read ND, Beckett A. 1996. Ascus and ascospore morphogenesis. Mycol
Res 100:1281–1314. https://doi.org/10.1016/S0953-7562(96)80057-8.
36. Freitag M, Hickey PC, Raju NB, Selker EU, Read ND. 2004. GFP as a tool to
analyze the organization, dynamics and function of nuclei and microtu-
bules in Neurospora crassa. Fungal Genet Biol 41:897–910. https://doi
.org/10.1016/j.fgb.2004.06.008.
37. Firman RC. 2018. Postmating sexual conflict and female control over fer-
tilization during gamete interaction. Ann N Y Acad Sci 1422:48–64.
https://doi.org/10.1111/nyas.13635.
38. Ryan FJ, Beadle GW, Tatum EL. 1943. The tube method of measuring the
growth rate of Neurospora. Am J Bot 30:784–799. https://doi.org/10
.2307/2437554.
39. Ainsworth GC. 1976. Introduction to the history of mycology. Cambridge
University Press, Cambridge, UK.
40. Ramos-Garcia SL, Roberson RW, Freitag M, Bartnicki-Garcia S, Mourino-
Perez RR. 2009. Cytoplasmic bulk flow propels nuclei in mature hyphae of
Neurospora crassa. Eukaryot Cell 8:1880–1890. https://doi.org/10.1128/EC
.00062-09.
41. Gladfelter A, Berman J. 2009. Dancing genomes: fungal nuclear position-
ing. Nat Rev Microbiol 7:875–886. https://doi.org/10.1038/nrmicro2249.
42. Roca MG, Kuo H-C, Lichius A, Freitag M, Read ND. 2010. Nuclear dynamics,
mitosis, and the cytoskeleton during the early stages of colony initiation
in Neurospora crassa. Eukaryot Cell 9:1171–1183. https://doi.org/10.1128/
EC.00329-09.
43. Mouriño-Pérez RR, Roberson RW, Bartnicki-García S. 2006. Microtubule
dynamics and organization during hyphal growth and branching in Neu-
rospora crassa. Fungal Genet Biol 43:389–400. https://doi.org/10.1016/j
.fgb.2005.10.007.
44. Lang C, Grava S, van den Hoorn T, Trimble R, Philippsen P, Jaspersen SL.
2010. Mobility, microtubule nucleation and structure of microtubule-
organizing centers in multinucleated hyphae of Ashbya gossypii. Mol Biol
Cell 21:18–28. https://doi.org/10.1091/mbc.e09-01-0063.
45. Nakamura K, Egashira T. 1961. Genetically mixed perithecia in Neuro-
spora. Nature 190:1129–1130. https://doi.org/10.1038/1901129a0.
46. Sansome ER. 1949. The use of heterokaryons to determine the origin of
the ascogeneous nuclei in Neurospora crassa. Genetica 24:59–64. https://
doi.org/10.1007/BF01487199.
47. Weijer J, Dowding ES. 1960. Nuclear exchange in a heterokaryon of Neuro-
spora crassa. Can J Genet Cytol 2:336–343. https://doi.org/10.1139/g60-036.
48. Cadot B, Gache V, Gomes ER. 2015. Moving and positioning the nucleus
in skeletal muscle: one step at a time. Nucleus 6:373–381. https://doi.org/
10.1080/19491034.2015.1090073.
49. Chalut KJ, Kulangara K, Giacomelli MG, Wax A, Leong KW. 2010. Deforma-
tion of stem cell nuclei by nanotopographical cues. Soft Matter 6:
1675–1681. https://doi.org/10.1039/B921206J.
50. Guilluy C, Osborne LD, Landeghem LV, Sharek L, Superfine R, Garcia-Mata
R, Burridge K. 2014. Isolated nuclei adapt to force and reveal a mechano-
transduction pathway in the nucleus. Nat Cell Biol 16:376–381. https://doi
.org/10.1038/ncb2927.
51. Dauer WT, Worman HJ. 2009. The nuclear envelope as a signaling node in
development and disease. Dev Cell 17:626–638. https://doi.org/10.1016/j
.devcel.2009.10.016.
52. Golloshi R, Martin RS, Das P, Raines TI, Thurston D, Freeman T, McCord RP.
2019. Constricted migration contributes to persistent 3D genome structure
changes associated with an invasive phenotype in melanoma cells. bioRxiv.
https://doi.org/10.1101/856583.
53. Freitag M, Selker EU. 2005. Expression and visualization of red fluorescent
protein (RFP) in Neurospora crassa. Fungal Genet Rep 52:14–17. https://doi
.org/10.4148/1941-4765.1124.
54. Margolin BS, Freitag M, Selker EU. 1997. Improved plasmids for gene tar-
geting at the his-3 locus of Neurospora crassa by electroporation. Fungal
Genet Rep 44:34–36. https://doi.org/10.4148/1941-4765.1281.
55. Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T,
Preibisch S, Rueden C, Saalfeld S, Schmid B, Tinevez J-Y, White DJ, Hartenstein
V, Eliceiri K, Tomancak P, Cardona A. 2012. Fiji: an open-source platform for bio-
logical-image analysis. Nat Methods 9:676–682. https://doi.org/10.1038/nmeth
.2019.
Brun et al.
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