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# Podospora anserina

Philippe Silar

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Philippe Silar. Podospora anserina. 2020, 978-2-9555841-2-5. hal-02475488

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Distributed under a Creative Commons Attribution - ShareAlike 4.0 International License
*Podospora anserina*

Philippe SILAR

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Podospora anserina
Acknowledgments.

Merci à Stéphan d'avoir été patient pendant que je rédigeais ce livre, souvent le matin très tôt ou le weekend ! Thanks to Professor Uwe Braun from the Martin Luther Universität at Halle-Wittenberg and Curator of Halle Herbarium (Germany) for both illustrations of type specimens. Merci à Pierre pour sa relecture !

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First edition : february 2020

ISBN : 978-2-9555841-2-5

EAN : 9782955584125
## SOMMAIRE

<table><tr><td>Foreword</td><td>1</td></tr><tr><td>Podospora anserina: a brief history</td><td>1</td></tr><tr><td>Part 1: Biology of Podospora anserina</td><td>3</td></tr><tr><td>Podospora anserina in the tree of life: classification of the species</td><td>4</td></tr><tr><td>Podospora anserina in its natural biotopes</td><td>26</td></tr><tr><td>Isolation, culture and preservation</td><td>29</td></tr><tr><td>Strains preservation</td><td>36</td></tr><tr><td>Methods for macromolecules extraction and genetic transformation</td><td>37</td></tr><tr><td>DNA Extraction</td><td>37</td></tr><tr><td>RNA Extraction</td><td>39</td></tr><tr><td>Protein Extraction</td><td>40</td></tr><tr><td>Protoplasts preparation</td><td>41</td></tr><tr><td>Genetic transformation</td><td>42</td></tr><tr><td>Gene deletion, single nucleotide change and at-will modification of the genome</td><td>44</td></tr><tr><td>Tools for expression analysis</td><td>49</td></tr><tr><td>Morphology and cytology</td><td>50</td></tr><tr><td>The mycelium</td><td>51</td></tr><tr><td>The gametes</td><td>55</td></tr><tr><td>The perithecium</td><td>57</td></tr><tr><td>The asci and the ascospores</td><td>60</td></tr><tr><td>The sexual cycle and genetical analysis</td><td>65</td></tr><tr><td>The lifecycle of P. anserina</td><td>65</td></tr><tr><td>Mutant generation</td><td>69</td></tr><tr><td>Genetical analyses: segregation, dominance/reluctivity and complementation</td><td>70</td></tr><tr><td>Parasexual cycle</td><td>74</td></tr><tr><td>Developmental genetics: grafting and genetic mosaics</td><td>74</td></tr><tr><td>Genes and genome</td><td>79</td></tr></table>
<table><tr><td>Part 2: <i>Podospora anserina</i> as a model for physiological and molecular analysis</td><td>89</td></tr><tr><td>Ascospore ejection and germination</td><td>91</td></tr><tr><td>Mycelium growth and development</td><td>94</td></tr><tr><td>Senescence and the Premature Death Syndrome</td><td>95</td></tr><tr><td>Crippled Growth and other phenotypic instabilities</td><td>103</td></tr><tr><td>Hyphal Interference and Vegetative/heterokaryon Incompatibility</td><td>111</td></tr><tr><td>Appressorium-like structure differentiation</td><td>119</td></tr><tr><td>Sexual reproduction</td><td>121</td></tr><tr><td>Mating and mating types</td><td>121</td></tr><tr><td>Fruiting body differentiation</td><td>125</td></tr><tr><td>Fruiting body production and repartition</td><td>132</td></tr><tr><td>Meiotic Drive Elements: Spore Killers</td><td>136</td></tr><tr><td>Natural products biosynthesis</td><td>140</td></tr><tr><td>Secondary metabolites</td><td>140</td></tr><tr><td>Enzymes</td><td>142</td></tr><tr><td>REFERENCES</td><td>145</td></tr><tr><td>Herbarium Specimen</td><td>145</td></tr><tr><td>Original Articles</td><td>145</td></tr><tr><td>Book Chapters</td><td>175</td></tr><tr><td>Dissertations</td><td>177</td></tr><tr><td>Conference Proceedings</td><td>181</td></tr><tr><td>Miscellaneous documents including edited thesis</td><td>182</td></tr><tr><td>Appendix 1: Original descriptions of <i>Podospora anserina</i></td><td>183</td></tr><tr><td>Appendix 2: Early papers on <i>Podospora anserina</i></td><td>191</td></tr></table>
# Foreword

In any good movie, the cast is a large part of its success. Of course, the stars playing the main characters are important. However, a good set of supporting roles is often equally crucial to fully bring to the fore all the qualities of the film. The science of Biology is a little bit like a movie. Indeed, it needs models to decipher the fundamental laws of the living. Some are superstars, such as mouse, drosophila, the bacterium *Escherichia coli* or the baker's yeast *Saccharomyces cerevisiae* and play foremost roles. However, other less-known models are sometime key actors in major discoveries. Examples are numerous: Mendel used peas to decipher laws of heredity and Hämmerling used Acetabularia to show that the cell's nucleus is the location where genetic information is stored...

For the filamentous fungi, the stars of the cast are *Neurospora crassa* and *Aspergillus nidulans*. The former has for example been instrumental in the discovery of the nature of the gene. Indeed, experiments on this model showed that "one gene codes for one enzyme", a paradigm which still holds for the most part. It is currently used in many labs to study many fundamental processes ranging from the circadian clock to epigenetic gene regulation. The second fungus is used to decipher many cellular phenomena ranging from the cell cycle to the role of the cytoskeleton. It has however been particularly important in the discovery of the parasexual cycle in eukaryotes in the 1950's. The supporting cast for filamentous fungi is too numerous to cite them all. *Cryptococcus neoformans* and *Magnaporthe grisea* are instrumental to study pathogenicity; *Coprinopsis cinerea* and *Sordaria macrospora* are used to study reproduction, to cite a few of them. This book will focus on one of the supporting cast, the "friendly mold" *Podospora anserina*. This fungus is used for over a hundred years in the laboratory to study processes as diverse as senescence, prions and sexual reproduction. Few reviews dealing with its biology are available. However, no large monography exists on this species. This book proposes to fill this void.
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**Podospora anserina: a brief history**

*Podospora anserina* is a filamentous fungus, i.e., a mold, used now for more than a century in several laboratories to study various biological processes ranging from those typical of fungi such as anastomoses and vegetative incompatibility (= heterokaryon incompatibility), hyphal interference, spore germination, mycelium degeneration and fruiting body formation, to those of general importance for eukaryotes such as sexual reproduction, Meiotic Drive Elements that cheat Mendel's laws, mitochondrial and peroxisomal physiology and those of general relevance in biology, e.g., senescence, cell differentiation, degeneration and death, respiratory metabolism, or structural and regulatory inheritance caused by prions and prion-like hereditary units.

Early researchers working with *P. anserina* were mostly concerned with the cytological descriptions of cellular phenomena and structures beyond those used to classify the fungus. Wolf<fnref n="1" /> in 1912 and Satina in 1916 dealt mostly with the description of the sexual process from the differentiation of the gametes to the production of ascospores, Ames in 1930, 1932 & 1934, Dowding in 1931 and Dodge

in 1936 with the breeding system, finally Buller in 1931 with hyphal anastomoses. Page analyzed the complete cycle of *P. anserina* and related species in 1939. It is however with the work of the French geneticist Georges Rizet (figure 1) that *P. anserina* was extensively used as a model. Rizet and its students gave the complete explanation of the pseudo-homothallic breeding system and complex nuclear behavior during sexual reproduction of the fungus, enabling them to perform elegant genetic analysis. They describe the vegetative

<figure><img src="image_6.png" /><figcaption>Figure 1. Georges Rizet.</figcaption></figure>

incompatibility (the “Barrage”phenomenon), the still mysterious Senescence process and the peculiar behavior of the S/s incompatibility system that we now know is due to a prion. They use the fungus to

<footnote n="1">¹ All references are given at the end of the book.</footnote>
study genetic recombination, gene structure, translation and mitochondrial physiology and evolution.
Along with Karl Esser, a German mycologist, who visited Rizet's lab during a postdoc in the 1950's, they
develop all the tools, which are now routinely used, including the growth and germination media as well
as the methods for macromolecule extractions and analyses. This culminated by the development of
genetic transformation procedures enabling the entry of *P. anserina* in the era of molecular genetics, by
permitting the identification of mutant genes and later on gene deletions and modifications. Finally, it is
Rizet's "scientific progeny" who established in 2008 the complete genomic sequence of the fungus.

Presently, the fungus is still studied in a few labs, mostly in France and Germany as expected
from its history. However, many publications also originate from the Netherlands, Mexico, Switzerland,
Taiwan... showing an ever increasing popularity! It truly deserves its nickname: the friendly mold...

<figure><img src="image_7.png" /><figcaption>**Figure 2.** *Podospora anserina*, the friendly mold.</figcaption></figure>

Picture courtesy Pierre Grognet.
# Part 1: Biology of *Podospora anserina*
*Podospora anserina* in the tree of life: classification of the species

When dealing with living organisms, it is always advisable to know their exact position in the tree of life as this greatly help to outline their main characteristics. *P. anserina*, also known as the "friendly mold", is thus a "mold" or a "filamentous fungus" (figure 3). This means that it belongs to the domain *Eukaryota* (figure 4), i.e., the major part of its genetic information is contained inside a nucleus, the remaining part being located inside the mitochondria. It is related to animals in the sense that like animals it belongs to the *Opisthokonta* superphylum (figure 4). Species of this superphylum share many characters, like the capacity to use extensively chitin as a coat material, to store carbon as glycogen and to use UGA as a tryptophan codon in the mitochondrial genetic code. Their name stems from the fact that they differentiate flagellated cells with a single posteriorly-orientated flagellum. However, *P. anserina* is a terrestrial fungus and, as such, it has altogether lost the capacity to differentiate the flagellum. *Opisthokonta* belongs to the subdomain *Amorphea*, which has been defined by molecular

<figure><img src="image_8.png" /><figcaption>**Figure 3** *Podospora anserina*. Left at high magnification (bar= 0.3 mm) and grown on toothpicks; Mycelium and fruiting bodies (the pear-shaped perithecia) are visible. Right: the friendly mold at low magnification (bar= 1 cm) and grown on a minimal medium (M2) Petri plate. The black dots are the perithecia.</figcaption></figure>
<figure><img src="image_9.png" /><figcaption>**Figure 4** *P. anserina* belongs to the *Eumycota*, or true fungi. In red, the lineage to which *P. anserina* belongs.</figcaption></figure>

phylogeny. Species from *Amorphea* share little characteristics, hence their name that means "formless". In *Opisthokonta*, two phylogenetic lineages are presently recognized: *Holozoa* that contains the *Metazoa* or animals, and *Holomycota* that contains the *Eumycota* or true fungi, to which *P. anserina* belongs. Fungi, and hence *P. anserina*, feeds by osmotrophy, i.e., they degrade their food outside the cells by secreting enzymes and then absorb the released molecules through transporters located in their plasma membrane. Species from the *Holozoa* lineage, and also those located at the base of the *Holomycota*, feed by phagotrophy or ingestion, i.e., ingest food as large particles and digest it inside the cells (or digestive
<figure><img src="image_10.png" /><figcaption>**Figure 5** Diversity of the Eumycota. In red, the lineage to which *P. anserina* belongs.</figcaption></figure>

track for animals). Other eukaryotic organisms feed by osmotrophy, the *Pseudomycota*. They look very much like *Eumycota*, but have evolved from completely different ancestors (figure 4).

*Eumycota* is a highly diverse group containing at least 14 phyla (figure 5). Its basal members, the “chytridiomycetes”, are still aquatic organisms that disperse thanks to a flagellum. However most species are terrestrial and have invaded nearly all biotopes. Some have even returned to an aquatic lifestyle, as whales and dolphins did. These terrestrial or formerly-terrestrial species all share the inability to produce a flagellum. The two major phyla that contain over 90% of the species of *Eumycota* are the *Ascomycota* and the *Basidiomycota*. They are related and able to produce dikaryotic cells (cells with two genetically-different nuclei) during an extended period of their life cycle, hence the name of the lineage that contains them; the *Dikarya* (figure 5). For most *Dikarya* species, the major part of their life is completed as a mycelium, a network of interconnected and elongated cells called hyphae. This mycelium is the stage of the life cycle during which the fungus feeds, i.e., the vegetative part of the life cycle. The major difference
<figure><img src="image_11.png" /><figcaption>**Figure 6** Diversity of the Ascomycota. In red, the lineage to which *P. anserina* belongs.</figcaption></figure>

between the *Basidiomycota* and the *Ascomycota* is the way they differentiate their sexual (meiotic) spores during the reproductive part of their lifecycle. In the *Basidiomycota*, spores are produced outside the mother cell or basidium that undergoes meiosis; they are called basidiospores. In the *Ascomycota*, they are formed inside the mother cell or ascus. They are then called ascospores. *P. anserina* belongs to the phylum of the *Ascomycota*, also known as the "ascomycetes" in the vernacular language. *P. anserina* differentiates a typical mycelium during its vegetative phase (figure 3) and produces archetypal asci during sexual reproduction (see those on figure 16).

*Ascomycota* are further subdivided into three subphyla (figure 6). The basal *Taphrinomycotina* and *Saccharomycotina* contain species unable to differentiate complex multicellular structures. They
<figure><img src="image_12.png" /><figcaption>**Figure 7** Diversity of the *Sordariomycetes*. In red, the lineage to which *P. anserina* belongs.</figcaption></figure>

often live as yeast or as mycelium in which communications between cells remains simple. *P. anserina* is a member of the third subphylum: *Pezizomycotina*. The species of *Pezizomycotina* often have complex life
cycles and behaviors, as we shall see for *P. anserina*. Their mycelium is highly interconnected, firstly by central pores that exist between two consecutive articles (i.e., the elongated cells that made up the hyphae), and secondly by the ability of hyphae to fuse by a process called anastomosis. Different types of hyphae can also be distinguished, primarily by their diameter. *Pezizomycotina* are often able to differentiate complex multicellular structures, especially during sexual reproduction. For example, *P. anserina* differentiate a fruiting body looking like a tiny pear that is called a perithecium (figure 3). *Pezizomycotina* fungi also disperse through asexual spores, often conidia. *P. anserina* differentiate conidia-looking cells that are used as male gamete for fertilization. They are called spermatia. As yet in the laboratory, germination of *P. anserina* spermatia is achieved with very low efficiency (in the range of one out of $10^6-10^7$), questioning their role as asexual dispersal unit. However, it cannot be ruled out that the proper conditions for their germination are still unknown.

Differentiating a perithecium during sexual reproduction is a general characteristic of species in the class *Sordariomycetes* (and also in the related class *Laboulbeniomycetes*) to which *P. anserina* belongs (figure 7). More than 10 000 species of *Sordariomycetes* have been described, but this number is likely largely underestimated. They live as saprobes (i.e., they live freely and feed on dead organic matter) or as parasitic or mutualistic associates of plants and animals. However, none appears to live as lichen or

<figure><img src="image_13.png" /><figcaption>**Figure 8** Diversity of *Sordariales*. In red, the lineage to which *P. anserina* belongs. The numbering is based on the one of Kruys *et al*. 2015.</figcaption></figure>
mycorrhiza, two lifestyles largely adopted by fungi. *P. anserina* lives primarily as a saprobe on herbivore dung and less frequently in soil, but it also seems to be able to associate with plants as an endophyte, i.e., it may also live within plants. Molecular phylogeny has recognized three major lineages of *Sordariomycetes*, as well as additional as-yet nameless basal groups (figure 7). Most species of *Xylariomycetidae* produce dark perithecia often embedded collectively within a stroma. Most species of *Hypocreomycetidae* produce brightly-colored perithecia and most species of *Sordariomycetidae*, to which *P. anserina* belongs, produced dark greenish ones. Molecular phylogeny has also helped to refine the classification of the various *Sordariomycetes* species into orders (figure 7). The order containing *P. anserina* is the *Sordariales*. Species of this order produce solitary perithecia that most often contain darkly-pigmented ascospores, as *P. anserina* does. *Sordariales* presently contains three families: *Sordariaceae*, *Chaetomiaceae* and *Lasiosphaeriaceae*. The definition of these families has greatly changed over time and was previously based on the fine structures of the fruiting bodies as well as the shape and ornamentation of the ascospores. The new data generated by the molecular phylogenies have shown that the *Lasiosphaeriaceae* is paraphyletic and four distinct phylogenetic groups can be identified. The monophyletic *Sordariaceae* and *Chaetomiaceae* are nested within these four lineages (figure 8). We are thus waiting for a complete reclassification of the order. This will entail name changes for three of the *Lasiosphaeriaceae* lineages... *P. anserina* is in the *Lasiosphaeriaceae* lineage IV that is more closely related to the *Chaetomiaceae*. Other well-known species of *Sordariales* (figure 9) are those of genera *Neurospora* and *Sordaria* belonging to the *Sordariaceae* and of the genus *Chaetomium* belonging to the

<figure><img src="image_14.png" /><figcaption>**Figure 9** Relatives of *P. anserina* from the order *Sordariales*. On the left, *Neurospora crassa* is known to produce large amounts of orange conidias. On the center, *Sordaria macrospora* produces typical glabrous perithecia. On the right, *Chaetomium globosum* differentiates very hairy fruiting bodies.</figcaption></figure>
<figure><img src="image_15.png" /><figcaption>**Figure 10** Main ascospore shapes in the *Lasiosphaeriaceae* IV. The size of the ascospores can be very different among the different species. Moreover, some can be decorated with gelatinous appendages not represented here.</figcaption></figure>

*Chaetomiaceae*. In addition to the famous *Neurospora crassa*, *Neurospora intermedia*, *Neurospora tetrasperma*, *Sordaria macrospora* and *Sordaria fimicola* are often used in the laboratories or in classrooms for genetic studies. *Chaetomium* species, especially *Chaetomium globosum*, are well adapted to grow on cellulose and are often responsible for the spoilage of books in humid libraries. Some *Chaetomium* species are responsible of very rare but often fatal mycosis in human. To give an insight into the biodiversity of the *Sordariales*, species in this order are as genetically diverse as the vertebrates. For example, the genetic divergence between *P. anserina* and *N. crassa* is at least as large as that between fishes and humans!

The traditional classification of the *Lasiosphaeriaceae* is based primarily on the form of the ascospores. Figure 10 depicts the shape of the ascospores of the species presently known in *Lasiosphaeriaceae* IV. Alas, this criterion turned out to be highly unreliable to trace the true relationship between species of *Lasiosphaeriaceae*. For example, most ascospores of *Lasiosphariaceae* IV are bicellular with one cell large and melanized and the other smaller and unpigmented. This latter cell may have undergone an apoptosis-like death. Nevertheless, *Arnium* ascospores are unicellular, showing that having bicellular ascospores is not a character shared by all species. Most genera of this family are thus polyphyletic. For instance, *Podospora* species are present in *Lasiosphariaceae* IV, but also in *Lasiosphariaceae* I, *Lasiosphariaceae* II and *Lasiosphariaceae* III! A paper by Miller & Hundorf in 2005 suggested that the peridium of the perithecium (= the envelop of the fruiting body that protects the asci)
<figure><img src="image_16.png" /><figcaption>**Figure 11** Known anamorphs in the *Lasiosphaeriaceae* IV.</figcaption></figure>

might be a better predictor of the true phylogeny. Unfortunately, too few species have been analyzed with regard to this character to know if it is actually able to predict relationships in the *Lasiosphaeriaceae*.

In addition to the species differentiating sexual structures (or teleomorph), several species of *Lasisopshaeriaceae* clade IV are only known through their asexual forms (or anamorph). These are known as *Cladorrhinum* (= *Bahupaathra*) or *Phialophora* (figure 11). Finally, it is most likely that many species of *Lasiosphaeriaceae* IV are presently unknown and those listed in figure 12 are likely to be only the “tip of the iceberg”.

To complicate the matter, few studies are devoted to the deciphering of the true phylogeny of the *Lasiosphaeriaceae*, as this family contains mostly saprobic species with inconspicuous life styles. Note that a fungus responsible for some mycetoma, *Madurella mycetomatis*, is a close relative, but it is not yet known whether this species belongs to *Lasiosphaeriaceae* IV or is more likely closely related to the *Chaetomiaceae*. Mycetomas caused by *M. mycetomatis* are rare but very debilitating and among the most dreadful diseases that one can catch! Presently, molecular phylogeny recognizes three subsets of species in *Lasiosphaeriaceae* IV (figure 12). In figure 12, the species of each subset are listed (mostly) alphabetically because their actual relationships are as yet unknown.

The two *Podospora* species most closely related to *P. anserina* are *P. setosa* and *P. austroamericana*. Both are also coprophilous fungi. *P. setosa* produces asci with 128 ascospores, *P. austroamericana* with eight ascospores, while *P. anserina* asci have four ascospores. *P. setosa* and *P. austroamericana* are homothallic, i.e., spores with a single nucleus will give rise to self-fertile thalli. *P.*
<figure><img src="image_17.png" /><figcaption>**Figure 12** The species most closely related to *P. anserina*. Precise phylogeny of these species is still not known. However, three subsets of species are defined in most analyses. One contains the "*Podospora anserina* species complex". Another contains *Podospora fimiseda*, the type species of the genus *Podospora*. The last one includes *Podospora australis*.</figcaption></figure>

*anserina* is pseudo-homothallic. Most of its ascospores give rise to self-fertile thalli. They in fact contain two kinds of sexually compatible nuclei and both are required for starting sexual reproduction. We shall come back to this point in the section dealing with *P. anserina* reproduction. It is not known whether these two *Podospora* are the actual closest relatives of *P. anserina*. Indeed, few molecular data are available for most of the species and the published phylogenies are poorly supported or partial (i.e., they only deal with a few species). Nevertheless, the best candidates appear to date to be *Cercophora samala* or *Zopfiella tetraspora*, because these two species have sequences of their Internal Transcribed Spacer (ITS) of the rDNA that are the closest to *P. anserina*. This is supported by their life style or their morphology (Table 1), the former being coprophilous and the latter producing four-spored asci. Note that *Zopfiella tetraspora* (also known as *Tripterospora tetraspora*) usually produces neckless perithecia (also
<table><caption>Table 1: some species related to P. anserina</caption><thead><tr><td>Species</td><td>habitat/life style</td><td>fruiting body</td><td>mating</td><td>Asci</td></tr></thead><tbody><tr><td>Apiosordaria backusii</td><td>soil</td><td>perithecium</td><td>homothallic</td><td>8-spored</td></tr><tr><td>Apiosordaria longicaudata</td><td>soil</td><td>perithecium</td><td>homothallic</td><td>4-spored</td></tr><tr><td>Apiosordaria tetraspora</td><td>soil and dung</td><td>perithecium</td><td>homothallic ?</td><td>4-spored</td></tr><tr><td>Apiosordaria verruculosa</td><td>soil, dung and endophyte</td><td>perithecium</td><td>pseudo-homothallic ?</td><td>4-spored</td></tr><tr><td>Apiosordaria yaeyamensis</td><td>soil</td><td>perithecium</td><td>homothallic ?</td><td>8-spored</td></tr><tr><td>Arnium arizonense</td><td>dung</td><td>perithecium</td><td>apomictic</td><td>4-spored</td></tr><tr><td>Cercophora samala</td><td>dung</td><td>perithecium</td><td>heterothallic</td><td>8-spored</td></tr><tr><td>Cercophora striata</td><td>decaying stems</td><td>perithecium</td><td>homothallic ?</td><td>8-spored</td></tr><tr><td>Cercophora squamulosa</td><td>aquatic decaying wood</td><td>perithecium</td><td>homothallic ?</td><td>8-spored</td></tr><tr><td>Cladorrhinum microsclerotigenum</td><td>endophyte of Musa sp.</td><td>anamorph</td><td>unknown</td><td>NA</td></tr><tr><td>Cladorrhinum phialophoroides</td><td>desert soil</td><td>anamorph</td><td>unknown</td><td>NA</td></tr><tr><td>Podospora austro-americana</td><td>dung and endophyte</td><td>perithecium</td><td>homothallic</td><td>8-spored</td></tr><tr><td>Podospora setosa</td><td>dung, soil and endophyte</td><td>perithecium</td><td>homothallic</td><td>128-spored</td></tr><tr><td>Triangularia batistae</td><td>soil and endophyte</td><td>perithecium</td><td>homothallic ?</td><td>8-spored</td></tr><tr><td>Zopfiella longicaudata</td><td>dung and soil</td><td>« cleistothecium »</td><td>homothallic ?</td><td>8-spored</td></tr><tr><td>Zopfiella tetraspora</td><td>dung and soil</td><td>« cleistothecium »</td><td>homothallic ?</td><td>4-spored</td></tr></tbody></table>

known as "cleistothecia", although this term is now reserved for species in the *Eurotiales*); yet an isolate was shown to produce both neck-endowed and neckless fruiting bodies... As seen in Table 1, many other potential applicants are possible. It is striking to see the diversity in the habitat/life style and developmental patterns of these species.

We are now finally reaching the species level in our journey through the classification of *P. anserina*. However, recent analyses of several strains of this "species" has reserved some surprises: *P. anserina* is a morpho-species - meaning that it has been defined by the morphology of its perithecia, asci and ascospores - that encompasses in fact at least seven species that appear to intercross rarely. All these species present different characteristics, including divergent genome sequences. Before going into the detail of each species, we need to go back to the traditional classification of *P. anserina*, which has seen battles of experts at to what is the proper name for this fungus!

Indeed, when one is looking in the fungal culture collections for strains of *P. anserina*, one is surprised to find that in some of them it is labelled as *Podospora pauciseta*. Moreover, in some early
<table><caption>Table 2: The different names of *P. anserina*</caption><thead><tr><td>Name</td><td>Reference</td></tr></thead><tbody><tr><td>Sphaeria pauciseta (Cesati)</td><td>Unknown author (1852) Botanische Zeitung 10: 285-288</td></tr><tr><td>Malinvernia anserina (Rabhenhorst)</td><td>Rabenhorst, L. (1857) Hedwigia 1: 116 - pl. 15 fig.4</td></tr><tr><td>Sphaeria anserina (Cesati)</td><td>cited as « in litt. » in Rabenhorst, L. (1857) Hedwigia 1: 116</td></tr><tr><td>Sordaria pauciseta (Cesati & De Notaris)</td><td>Cesati, V & De Notaris, G (1853) Schema di classificazione degli sferiacei italici aschigeri pp 51-53</td></tr><tr><td>Sordaria anserina (Rabhenhorst) Winter</td><td>Winter G. (1873) Botanische Zeitung 31: 481-485</td></tr><tr><td>Hypocopra anserina (Cesati)</td><td>cited as « in litt. » in Sacchardo P.A. (1882) A Sylloge fungorum omnium hucusque cognitorum. 1: 238</td></tr><tr><td>Hypocopra erecta (Spegazzini)</td><td>Spegazzini C. (1880) An. Soc. Cient. Argentina 10: 5-33</td></tr><tr><td>Podospora anserina (Rabhenhorst) Niessl</td><td>Niessl G. (1883) Hedwigia 22: 153-156</td></tr><tr><td>Sordaria Penicillata (Ellis & Everhart)</td><td>Ellis, J. B. and B. M. Everhart (1888). The Journal of Mycology 4(8): 73-82.</td></tr><tr><td>Pleurage anserina (Rabhenhorst) Kuntz</td><td>Kuntze, o. (1898). Revisio generum plantarum. 3(2): 504-505.</td></tr><tr><td>Sordaria communis var. tetraspora (Spegazzini)</td><td>Spegazzini, C. (1899). An. Mus. Nac. Hist. Nat. Buenos Aires Ser. 2 6: 289-365.</td></tr><tr><td>Podospora pauciseta (Cesati) Traverso</td><td>Traverso, J. B. (1905) Flora Italica Cryptogama ParsI: Podospora pauciseta. 1(2): 431-432</td></tr><tr><td>Bombardia anserina (Rabhenhorst) Migula</td><td>Migula, W. (1913) Thome's Kryptogamic Flora. 10: 123-129.</td></tr><tr><td>Schizothecium anserinum (Rabhenhorst) Bessey</td><td>Bessey, E. A. (1950). Morphology and Taxonomy of Fungi. pp 264-265</td></tr></tbody></table>

papers, this fungus was called *Pleurage anserina*. *Pleurage anserina* is only one of the names that this fungus has been designated and a full list is given in Table 2². This proliferation of names stems from the fact that different authors classified the fungus under different names, for taxonomic purposes. Indeed, the genus is supposed to reflect on the relationships between close species. Depending on the characters used to regroup species, as well as the concept of “close” by the mycologist that has first identified the species, a fungus ends up in an already-known genus or in a new one. The “type” for the new fungus should at the same time be deposited in a herbarium or a culture collection for future analyses. This is called an exsiccata in the case of dried specimen kept in herbaria and it has a voucher for further reference. Then, as knowledge progressed, and as new species are identified, another taxonomist may reexamine the fungus and its name may change because new genera are created to accommodate growing numbers of species (or because this new specialist deemed his/her own set of characters to be important for classification!). Normally, during these transfers between genera the species epithet should

<footnote n="2">² Table 2 may be incomplete, as some authors, such as F. Doveri in “Fungi Fimicoli Italici”, list more synonyms, but direct consultation of the cited literature does not permit to conclude that the described species is indeed our friendly mold.</footnote>
42. *Sph. pauciseta* Ces. mss. *Pyrenia minuta*, sparsa, primitus solo verice setis paucis fasciculatis arrectis comato conspicua, dein emersa, ostiolo tandem denndato, deciduo (?), papillaeformi. Nuclens ascigerns, ascis paraphyses superantibus; sporidia 1-seriata ovalia, simplicia.

42. *Sph. pauciseta* Ces. mss. Perithecium small, sparse, at first like a wart with few coarse upright hair, then emerging, finally with an ostiole naked and deciduous (?), forming a papilla. Formation of asci in the center, asci taking over the paraphyses; spores uniseriate, oval, simple.

**Figure 13** First description of *P. anserina* as *Sphaeria pauciseta* in the Botanische Zeitung in 1852 vol. 10 pp 285-288.

not change (although it may be corrected to comply with the Latin terminology). Another source of names is that a species may be identified as new, while it was already known. When this is realized, the two names are synonymized and the first one should be used. All of this participate to a proliferation of names and add to the confusion. Note that this is not restricted to *P. anserina*, but is common for many fungi, especially when they are molds with tiny fruiting bodies...

So why two species epithets for our friendly mold? Well, the proper name being normally the first one given, it should be *P. pauciseta* because the fungus appears to have been first formally described in 1852 as a "*Sphaeria pauciseta*" by an italian botanist called Vincenzo de Cesati (1806-1883). It was deposited in the "Klotsch herbarium mycologicum" under the number n° 1642. However as seen on figure 13, the description given in the Botanische Zeitung of this new species is rather limited (especially, it has no associated iconography) and the description from the herbarium associated with the original exsiccata is identical<fnref n="3" />. Note that in the description the number of ascospores in asci is not given, nor are the sizes of perithecia, asci and ascospores. There is thus no way to know based on this description whether this fungus is actually *P. anserina* with its typical four-spored asci. Many species of *Sordariales* may fit the description of the Botanische Zeitung. There is for example *Podospora austroamericana* having asci with eight ascospores that may also fit to the description for "*Sphaeria pauciseta*". This description was nonetheless validated by Giovanni Battista Traverso (1878-1955), an italian mycologist in 1907 in his "Flora Italica Cryptogama", this time with an associated drawing (figure 14). The depicted species seems to be indeed our friendly mold, hence, the name validated by the taxonomists, "*Podospora pauciseta* (Ces.) Trav." that is used in some papers and culture collections.

<footnote n="3">³ Pictures of the original exsiccatas, as well as old publications dealing with the fungus, are appended in the annexes at the end of the book.</footnote>
<figure><img src="image_18.png" /><figcaption>**Figure 14** Illustration of *P. pauciseta* by Traverso in his "Flora Italica Cryptogama". Note the similarity of these drawings with those of Griffiths published six years earlier (figure 17)...</figcaption></figure>

The second historical and formal mention of the fungus appears to be by the German mycologist Gottlob Ludwig Rabenhorst (1806-1881) as "Malinvernia anserina" in the first issue of the journal Hedwigia. The description is also rather scant (figure 15), but is associated with some drawings, most likely the first ones for our friendly mold. In fact, "Malinvernia anserina" is only described in the legend of a figure! The type for this "Malinvernia anserina" was deposited under n° 526 in the "Klotschi herbarium vivum mycologicum sistens fungorum per totem Germaniam crescentum collectionem perfectuam, ed. II". There, the description is more extensive. Many mycologists consider this to be the first accurate description of *P. anserina* and hence prefer to use "anserina" as the species epithet, especially given the
<figure><img src="image_19.png" /><figcaption>**Figure 15** Second description of *P. anserina* as *Malinvernia anserina* in the first volume of Hedwigia p 116 Fig. 4 of plate 15.</figcaption></figure>

doubts that shroud the first description as *Sphaeria pauciseta* and the fact that Cesati also described a *"Sphaeria anserina"* and a *"Hypocopra anserina"* in earlier letters (see in litt. in Table 2). Unfortunately, I have not been able to find these letters nor their date of writing, to ascertain whether Cesati was indeed referring to the same species.

All this confusion regarding the proper name was already noted by the American mycologist George Francis Atkinson (1854-1918) in a footnote of a paper published in 1912 by another American mycologist, Frederick Adolph Wolf (1885-1975). Atkinson recommended using the name *P. anserina*... In fact, we may never know which description for the fungus is the good one, even if we go back to the herbarium specimens. Indeed, at that time pure cultures were rare and descriptions often relied on samples collected from the wild and not on strains isolated in pure cultures. The original specimens for *P*.
<figure><img src="image_20.png" /><figcaption>**Figure 16** *Podospora tetraspora*. This four-spored species may be the one actually described as *Malinvernia anserina*. It produces slender perithecia with a differently-shaped neck, as well as smaller ascospores. The bottom pictures are from *P. anserina* taken at the same magnification for comparison. From left to right, bar= 250 µm, 250 µm & 100 µm.</figcaption></figure>

*pauciseta* and *P. anserina* consist in dried dung with potentially more than one fungal species on it! Accordingly, in his thesis “Nordic Sordariaceae S. Lat.” the Swedish mycologist Nils Lundqvist mentions that he was not able to find *P. pauciseta* in the “authentic” collections he examined. Intriguingly, it seems to me that the dimensions of the spores given by Rabenhorst, the long appendages on the neck of the fruiting bodies as well as the presence of a bubble in the center of the spores (discernible in figure 15), fit more with *P. tetraspora* than with *P. anserina...* This *Podospora* species looks very much like *P. anserina* (figure 16). It has similar-sized and -looking perithecia with four-spored asci, but these are smaller than in *P. anserina*. This species belongs to *Lasiosphaericeae* clade I. This casts strong doubts on the original description of *Malinvernia anserina* being that of our friendly mold. Along this line, some authors such as Mirza & Cain in their “Revision of the Genus *Podospora*” in 1969 even state that it is doubtful that *P. pauciseta* and *P. anserina* are the same species! So it is possible that neither “*pauciseta*” nor “*anserina*” should be the proper species epithet...

The quality of the microscope in the mid-19th century likely prevented a more accurate description of our friendly mold by Rabenhorst (the drawings of figure 15 show many inaccuracies in the appendage of the spore and the neck of the perithecia even if the represented species is *P. tetraspora*
<figure><img src="image_21.png" /><figcaption>**Figure 17** Two early drawings of *P. anserina*. Left (fig. 15-21), by E.C. Hansen in 1876 under the name *Sordaria anserina*; Right (fig. 4-6) by D. Griffiths in 1901 under the name *Pleurage anserina*. Compare the right drawings with those of Traverso (figure 14).</figcaption></figure>

and not *P. anserina*). Though, improvement in microscope quality rapidly permitted to obtain better drawings for the fungus. Figure 17 gives two of them showing the actual *P. anserina*. The first one by the Danish mycologist Emil Christian Hansen (1842-1909) in his "Champignons Stercoraires du Danemark" published in 1876 and the second one by the American mycologist David Griffiths (1867-1935) in his "North American *Sordariaceae*" published in 1901. It is likely that Traverso (figure 14) got some inspiration from the earlier drawings from Griffiths (figure 17)...

At the beginning of the XXth century, confusion was already high regarding *P. anserina*, when in 1937 a Ukrainian mycologist, M. Milovtzova, described a new species closely related to *P. anserina* and
named it *Podospora comata* (figure 18). This species was described as having slightly smaller ascospores and perithecia. On the provided drawing, the neck of the peritheium lacks the small brush of hair at the base of the neck that is characteristic of *P. anserina*. This species was subsequently considered either a true species or only a "minute" form of *P. anserina*, depending on the mycologist...

<figure><img src="image_22.png" /><figcaption>**Figure 18** *Podospora comata*, a new species described by M. Milovtzova.</figcaption></figure>

This species is described as having smaller ascospores and perithecia. The drawing on the left of the peritheium does not mention the presence of the small brush of erected hair typical of *P. anserina*. This species was considered either as a true species or as a minute form of *P. anserina*.

Owing to the doubts concerning the original descriptions of *P. pauciseta* and *P. anserina*, as well as the uncertain status of *P. comata*, we reexamined in my lab many strains conserved under these names in culture collections. Thanks to the molecular tools now available, we were able to show that the strains stored in the culture collection under the three names belong in fact to seven *bona fide* species, i.e., populations that likely rarely interbreed in the wild, although they can mate with each other in the laboratory. This means that they accumulated throughout evolutionary times many differences or polymorphisms in their genome. Quantification of the differences between the seven species shows that their genomes differ in average by 1-4% at the nucleotide level, a difference similar to the one between the genomes of human and chimpanzee. Moreover, many genes present in one species, may be missing in the others. We identified for example genes encoding a laccase, a histone or a cytochrome P450 as
<figure><img src="image_23.png" /><figcaption>**Figure 19** Perithecium production patterns of the species from the *P. anserina* species complex are species-specific. The medium is the minimal M2 medium with 4g/L of potato dextrins. On other media, the strains will exhibit different patterns.</figcaption></figure>

being present in the strain named T and absent in our reference strain named S. Importantly, phenotypic
analysis showed that the criteria traditionally used to differentiate *P. comata* from *P. anserina* were not
valid. For example, strains belonging to the same species could be labelled under different names in the
collections and reciprocally strains having different names could belong to the same species! On the
contrary, careful analyses showed that the seven species could be differentiated by the way they
produced fruiting bodies on several media differing by the carbon source (figure 19). Some also exhibited
typical phenomenon not displayed by the others (Table 3). We were able thus to name these seven
species and provided new types for them. We kept the three already used species epithets (*anserina*,
*pauciseta* and *comata*) for three species that we formally redefined and proposed four new names
(bellae-mahoneyi, pseudoanserina, pseudocomata and pseudopauciseta). Of course, we chose to give the
name *P. anserina* to the species to which our major working strain belong (strain S or BIG S). Most work
on *P. anserina* has been made with this strain or with strain s (small-s), which fortunately also belongs to
the *P. anserina* species as newly redefined. We designated as belonging to the redefined *P. comata*, the
<table><caption>Table 3: some features of the species from the *P. anserina* species complex.</caption><thead><tr><th></th><th>Hyphal Interference<sup>1</sup></th><th>Crippled Growth<sup>1</sup></th><th>ring of perithecia<sup>1</sup></th><th>Senescence<sup>1</sup></th></tr></thead><tbody><tr><td><i>P. anserina</i></td><td>efficient</td><td>yes</td><td>yes</td><td>yes</td></tr><tr><td><i>P. pauciseta</i></td><td>inefficient</td><td>no</td><td>no</td><td>yes</td></tr><tr><td><i>P. comata</i></td><td>inefficient</td><td>no</td><td>no</td><td>yes</td></tr><tr><td><i>P. bellae-mahoneyi</i></td><td>inefficient</td><td>no</td><td>no</td><td>yes</td></tr><tr><td><i>P. pseudoanserina</i></td><td>inefficient</td><td>no</td><td>no</td><td>yes</td></tr><tr><td><i>P. pseudocomata</i></td><td>inefficient</td><td>no</td><td>yes</td><td>yes</td></tr><tr><td><i>P. pseudopauciseta</i></td><td>inefficient</td><td>no</td><td>no</td><td>yes</td></tr></tbody></table>

aforementioned strain called T. This strain has previously been used in few molecular studies under this name. It is the only publicly known cultivated isolate for this species. Fortunately, strain T produce smaller ascospores and perithecia, as did the original isolate described by Milovtzova as a new species. We chose *P. pauciseta* for a group of three strains, since this will limit name change in the culture collections. These have not been yet used for genetical or biochemical studies, nor have the strains belonging to the four new species. Nonetheless, the seven species share common features. Their ascospores germinate with the same modalities. Their mycelium grows, differentiates aerial hyphae and

<table><tr><td>Eukaryota</td><td>Domain</td></tr><tr><td>Amorphea</td><td></td></tr><tr><td>Opisthokonta</td><td></td></tr><tr><td>Holomycota</td><td></td></tr><tr><td>Eumycota</td><td>Kingdom</td></tr><tr><td>Dikarya</td><td></td></tr><tr><td>Ascomycota</td><td>Phyllum</td></tr><tr><td>Saccharomyceta</td><td></td></tr><tr><td>Pezizomycotina</td><td>subphyllum</td></tr><tr><td>Leotiomyceta</td><td></td></tr><tr><td>Sordariomyceta</td><td></td></tr><tr><td>Sordariomycetes</td><td></td></tr><tr><td>Sordariomycetidae</td><td></td></tr><tr><td>Sordariales</td><td></td></tr><tr><td>Lasiosphaeriaceae IV</td><td></td></tr><tr><td>Podospora anserina species complex</td><td></td></tr><tr><td>Podospora anserina</td><td></td></tr></table>

**Figure 20** Current Identity card of *P. anserina*. This card has been established thanks to molecular phylogenetic analyses. Except for potential modifications in the names of the family and genus, pending a revision of the *Lasiosphaeriaceae*, this identity card should now not change. Prior, *P. anserina* was included in the now defunct *Pyrenomycetes* class, *Sphaeriales* order and *Sordariaceae sensus Lato* family.

<footnote n="1">¹These phenomena will be described in the chapter “Physiological and molecular analysis: deciphering developmental pathways”.</footnote>
accumulates pigment in a similar fashion, even if minor differences between the strains can easily be
spotted. They all undergo senescence (Table 3).

P. anserina, as usually known, is thus a species-complex with at least seven members that cannot be distinguished by simple visual inspection. It is likely that several more species belonging to the complex as yet not isolated exist in the wild. The now newly-redefined P. anserina is only one of the species of the complex. This book will be focused on this particular one with some references to P. comata. Its complete identity card is given in figure 20. To summarize, P. anserina is a filamentous fungus from the Pezizomycotina subphylum of the Ascomycota. As such, it has a lifecycle typical of this group of fungi, including the ability to differentiate a multicellular fruiting body. This fruiting body is a perithecium having a greenish color, typical of the Sordariales.

In a turn of fate, as this book was in its final stage of writing, a paper starting to partially revise
the phylogeny of *Lasiosphaeriaceae* was published (in august 2019 by Wang et al. in *Studies in Mycology*)
in which the name of *Podospora anserina* was changed into "*Triangularia anserina* (Rabenh.) X. Wei
Wang & Houbraken", and all the species of the complex had accordingly the name of their genus changed
to *Triangularia*. A name for the *Lasiosphaeriaceae* Clade IV was also given: *Podosporaceae*. While it is
most likely that the name *Podosporaceae* will stick for *Lasiosphaeriaceae* Clade IV, I think it is highly
unlikely that the change to *Triangularia* will be adopted. Indeed, although it would be the correct way to
name the friendly mold, two reasons militate against its usage. Firstly, researcher working on ageing,
prions, sexual development, etc., especially those not working on fungi will not understand the need for a
change and will thus not use the new name. This is especially true because several hundreds of papers
have already been published on the friendly mold with its *P. anserina* name, and this is not counting the
thousands of paper citing studies with this fungus! Changing the name of the fungus will thus only lead to
great confusion. Note that very few papers have been published (I am aware of only two) on *Podospora
fimiseda* the type species of the genus, apart from the purely taxonomic ones. According to me it would
therefore be wise to maintain for all eternity the name of friendly mold as *Podospora anserina*. This can
be done only by changing the type species for *Podospora* to *P. anserina*; however this is not an easy task
as taxonomist tends to be very very conservative... Note that intense battles other the names of famous
fungi (such as *Aspergilli*) have lately been won by changing the International Code of Botanical
Nomenclature. Facilitating the change of genus types for fungi such as *Podospora*, whose origin is
obscure since the herbarium types for the species although available are not useful would be
appreciated... Ironically, the taxonomic origin of *Podospora fimiseda* is as mysterious as the one of *P. anserina*, if not more!

A second reason for not adopting the name is that it is most likely that, according to the rule of nomenclature, it will change again shortly! Indeed, the genus adopted for the new name is highly diverse. *P. anserina* is in a lineage different from *Triangularia bambusae*, the type species for *Triangularia*. Hence, once additional species are identified of *Podosporaceae* and their phylogeny sorted out, it is most likely that *Triangularia* will be split into many genera! Note that as explained above *P. anserina* has already suffered many battles upon its naming. In the end, it appears that the name *Podospora anserina* has always prevailed...

Therefore, whatever the fate of the new naming to *Triangularia*, I have decided to conserve the name under which the friendly mold is known and that is *Podospora anserina*!!
*Podospora anserina* in its natural biotopes

Many fungal species have evolved through natural selection life strategies that permit them to use dead plant material as carbon and energy sources. These fungi produce and export outside the cells many enzymes that allow them to degrade plant polymers (like cellulose or lignin). Released nutrients are then transported into the fungal cell by very efficient transport systems. Such nutrition strategy is called saprotrophy and the fungi are said to be saprotrophic, saprophytic or saprobic. *P. anserina* is one such a saprotrophic fungus that has specialized to retrieve its nutrients from materials that have not been completely digested by herbivorous vertebrates, i.e., their dung. About 2000 fungal species are known to inhabit dung including many *Podospora* species. They are called coprophilous, coprophilic or fimicolous.

Coprophilous fungi fructify sequentially in a fashion that recapitulate what we know about fungal evolution. The first ones to be observed are basal fungi such as species from genera *Mucor* or *Pilobolus*, then basal *Ascomycota* from the class *Pezizomycetes* such as species from genera *Ascobolus* and *Saccobolus*. These are followed by species from the class *Sordariomycetes* including those from genera *Sordaria* and *Podospora* that appear just before the final *Basidiomycota* from genera *Coprinopsis*, *Coprinus* or *Cyathus* among others. This succession experiences nonetheless many exceptions and it is not rare for example that fast-growing *Coprinus* or *Coprinopsis* appear early and prevent appearance of other fungi. It is likely that all fungi are inoculated in the dung as spores stuck on the plants ingested by the animals. Spores are triggered to germinate while passing through the digestive track. All the coprophilous fungi likely start thus to grow at about the same time. The observed succession is in fact the complex result of interactions between growth speeds of the mycelia and timing of differentiation of the sporophores. Both are modulated upon the ability of each species to use more or less hard to digest plant remnants and their ability to eliminate the bacterial and fungal competitions.

*P. anserina* is usually one of the last species to fructify. Although very rapid on synthetic medium or sterile dung (the complete cycle is then completed in one week), when in competition with other microorganisms, *P. anserina* takes more time to fructify (about two to three weeks). Its growth speed of about 7 mm/day is slower for example than that of *Sordaria macrospora* which is of 2-3 cm/day.
<figure><img src="image_24.png" /><figcaption>**Figure 21** *P. anserina* perithecia on its natural biotope: dung. The horse dung used here to cultivate the fungus is composed of partially digested plant debris, which can be further broken down thanks to the numerous enzymes encoded in the genome of *P. anserina*. Bar=0.5 mm.</figcaption></figure>

However, the genome of *P. anserina* contains more genes coding for enzymes enabling to cope with lignocellulose and the fungus exhibits hyphal interference towards some fungi, while *S. macrospora* does not. Hyphal interference is a mechanism whereby hyphae are able to kill hyphae from other species upon contacting them. Although the fruiting bodies of the two fungi appear at roughly the same time on dung, to do so *S. macrospora* rely mostly on its fast growth and utilize easy to reach cellulose, while *P. anserina* appears to count on its abilities to extract nutrient from harder to digest plant debris and to kill competition.

Investigation of *P. anserina* in the wild has presently been made only by visual determination of the perithecia (figure 21), with isolation in pure cultures and molecular determination only in few cases.
Therefore, there is no way to know whether the observed specimens belonged actually to *P. anserina* or
to one of the others from the complex. Inventories of fungi growing on dung have shown that these are
frequently found on many kinds of dung from birds and mammals originating from all regions of the
world. The exact geographic distribution of *P. anserina* is unknown. We know that the fungus is
commonly found in Western Europe during summer, because all strains of *P. anserina* but one hosted in
culture collections come from this region. The only exception originates from Ontario in Canada. The
strains from culture collections isolated from other regions of the world often belong to the other species
of the complex, tentatively suggesting that there is a geographical structuration of the complex. Each
species would have evolved to adapt to the faunas/floras present in each region. This would nicely fit
with the fact that they seem to utilize carbons sources differently (see previous chapter, figure 19). *P. anserina* would thus be the one adapted to mild climates. However, most *P. anserina* strains were
isolated from domestic horse and cow dung and it is possible that it cohabits with other species more
adapted to dung from other herbivores including birds like geese or smaller mammals like hares, rabbits
or even mice.

In addition to being collected from dung, species from the *P. anserina* complex have also been
isolated from a decaying Chinese mat, soil and living plants. Its presence in soil and as a plant endophyte
is confirmed by metagenomic data, since its DNA is sequenced along with that of related species when
analyzing various plants and soils. At the present time, it is not clear if these alternate ecological niches
are part of the normal cycle of the fungus or are occupied accidentally.

Overall, we know little about *P. anserina* in its natural biotopes, unlike for example *Neurospora spp.* for which extensive data regarding strain variations exist. To better understand *P. anserina*, and the other species from the complex, we now need to investigate its population structure thanks to molecular technologies. Extensive analyses of many isolates collected around the world will permit to identify more species and to understand if there is a geographical structuration of the populations related to the particular faunas and floras present in the ecosystems. Another needed line of investigation is the catalogue of all the natural biotope the fungus is able to invade.
Isolation, culture and preservation

In this section, I will describe how to collect, grow and preserve *P. anserina* and the other species of the complex, since they all behave similarly. The recipes for the media are the optimal ones and I will not in this section examine how modifications of the media impact on the fungus. This will be dealt in the appropriate sections regarding the modalities of germination growth and reproduction. The toolkit required to isolate and cultivate the fungus is rather simple. In addition to a 10-40 X binocular microscope, it should include tools to manipulate ascospores and mycelia (figure 22). The fungus can be grown at room temperature, but a temperature-controlled chamber should ensure reproducible result. Ideally, it should have also controlled humidity and light. Indeed, optimal growth conditions are 27°C, 70-80% humidity (to prevent desiccation of the Petri plates) and constant illumination or 12 hours alternation of light and dark (to allow for perithecium production). There is no need for a sterilized chamber and all manipulation can be performed on the bench.

If you wish to recover *P. anserina* strains from nature in order to grow them in the lab, you need to fetch dung from the fields. Horse dung seems rather efficient, but any kind of herbivore dropping

<figure><img src="image_25.png" /><figcaption>pen point holder and nib</figcaption></figure>

<figure><img src="image_26.png" /><figcaption>mounted needle</figcaption></figure>

<figure><img src="image_27.png" /><figcaption>alcohol burner</figcaption></figure>

<figure><img src="image_28.png" /></figure>

**Figure 22** The basic tool kit to work with *P. anserina*. Pen point holders are used to slice explants of mycelium from jellified growth plates. The ones with replaceable nibs are ideal. Mounted needles or "pique-huile" are used to collect fruiting bodies and ascospores. These tools can be sterilized with an alcohol burner.
<figure><img src="image_29.png" /><figcaption>**Figure 23** Humid chamber and projection plates are ideal to collect new P. anserina strain from the wild.</figcaption></figure>

should do; the fungus has seemingly been first seen on pig and goose dung! Fresh dung is preferable since in old ones it may have already fructified. Incubate the dung in a closed and humid container as to make a humid chamber and let the various fungi grow. Perithecia can be easily spotted on the dung after 10-20 days of incubation and ascospores can be recovered on a projection plate as described in figure 23. Recipe for the projection plate is given in the "projection plate" box. On wild dung, many fungi should grow, some having morphologies quite similar to our friendly mold. I find it easier to recognize the asci of P. anserina once expelled on the projection plate than the perithecia on dung; those look quite similar to those of many other species.

Perithecia may be collected with a mounted needle or a "pique-huile" and transported onto a fresh projection plate (see movie n°1). After bursting the perithecium, ascospores may be collected also with a mounted needle and transported onto germination medium. Antibiotics could be added to the germination plates to prevent bacterial contamination. To burst the perithecia, simply squeeze them with a thin tweezer or between two mounted needles or "pique-huiles". Rosettes of asci are liberated. Individual asci can then be gently probed with a mounted needle as to break them apart. Usually the four ascospores stick together thanks to a small rope connecting them. The trick to collect the ascospores is then to break apart this rope. This demands some skill that one usually masters in two or three sessions of ascospore collection (movie n°2). Individual ascospore stick reversibly to the mounted needle and can be deposited at will on fresh media. Note that the needle must be sterile and this is achieved by flaming
**Projection plates:** Also called “agar covers”, these are used to recover ascospores that are ejected outside the perithecia. Ascospores on these covers can resist for a few weeks at 4°C before dying. Projection plates are made by pouring a thin layer of melted agar (20 g/L) supplemented with NaCl (10 g/L) in a Petri plate. Autoclave for 20 min at 120°C and pour (12 mL for an Ø=8 cm plate) very hot as to make the agar very compact.

the tip. Once flamed, the tip is very sticky and should be re-humidified by cooling it in the jellified medium of the projection plates, otherwise ascospores will stick irreversibly on it. Note that starting directly from the projection plates has three advantages. Firstly *P. anserina* asci are easily recognized, since the four ascospores often stick together until landing on the projection plates. Their morphology is quite distinctive (see next chapter). Secondly, bacterial contamination is less frequent. Finally, breaking apart the rope is somewhat easier, especially if the projection plates were stored at 4°C for a few days.

Once ascospores are recovered, they can be transported onto Germination plates containing G medium. I usually put 12-16 ascospores per plate (figure 24). Germination will proceed overnight at temperature ranging from 18°C to 35°C with nearly 100% efficiency. We routinely use 27°C as it appears to be the optimal growth temperature. One day after collection, the germination thalli are often barely visible to the naked eye, but two days after germination they should have a diameter of over one cm (figure 24). Alternatively, ascospores may be collected on M2 medium. A heat shock is then required to awaken the ascospores (25-30 minutes at 65°C works best). With this procedure up to 90% of the ascospores may germinate. However, lower percentages are often obtained.

<figure><img src="image_30.png" /><figcaption>**Figure 24** Typical figures of germination plates. Here, 4 asci (=16 ascospores) have been isolated. Left: after ascospore collection, arrows point the locations where the ascospores were placed, middle: one day after colection, germination thalli are still not visible with the naked eye, right two days after collection.</figcaption></figure>
**Germination plates:** These plates containing G medium are mainly used for ascospore germination. G medium allows the germination of nearly 100 % of wild-type ascospores. On this medium, the thalli never produce pigment (figure 25). Composition of G medium is ammonium acetate (4.4 g/L or 10 mL from a 440 g/L solution), bactopeptone (15 g/L) and agar (13 g/L). Autoclave (20 min at 120°C) and pour 23 mL per Ø=8 cm plate. Variations in germination efficiency and size of germination thalli are observed depending upon the age and brand of the components used to make G medium, especially water. Fresh bacto-peptone is better; if too old the germination percentage may drop. This can be corrected by adding yeast extract (5 g/L). This product should also be rather fresh as it inhibits growth when too old.

Like all the filamentous fungi, *P. anserina* will produce a mycelium on its growth media. Germination thalli are thus typical mycelia. It is possible to cut them into small pieces (to do so we use a pen point holder whose nib has been sterilized with an alcohol burner). Each piece should regenerate a new thallus when inoculated onto fresh growth medium. At the macroscopic level, the mycelium grows as a disk. However, it can display many different aspects, depending upon the growth medium (figure 25). Some media permit the differentiation of numerous aerial hyphae and the mycelium looks velvety. Others prevent the growth of such hyphae and the mycelium is smooth. Color also may vary. Most of the time it is greenish but it may also be brownish or pigments may be absent.

We routinely use the M2 Medium at 27°C in the presence of light to grow our friendly mold. Crosses, stocks and most phenotypic tests are made on it. It permits the mycelium to grow at about 7mm per day. In stationary phase (figure 25), the mycelium develops aerial hyphae, presents a greenish pigmentation and, if *mat+/mat-* heterokaryotic (see chapter “The sexual cycle and genetical analysis”), it should undergo sexual reproduction in a week with four days between fertilization and ejection of the ascospores out of the perithecium.

Another frequently used medium is Corn Meal medium (that we often call MR). It is a semi-defined medium that we use less and less because M2 is easier to prepare and often satisfy most needs. On this medium, the mycelium develops no or few aerial hyphae (figure 25) and pigmentation is variable, but is often more pronounced that on M2. Fertility is lower than on M2 and the sexual cycle takes longer as it is completed in about ten days. An advantage of this medium is that it contains almost all amino acids, bases, vitamins, etc. Hence, most auxotrophic mutants grow on this medium. The mycelium will also not penetrate cellophane and can thus be scraped more easily than when grown on M2. Cellophane
<figure><img src="image_31.png" /><figcaption>**Figure 25** *P. anserina* mycelium morphology on three commonly-used media. The strains inoculated on the plate are homokaryons that do not differentiate fruiting bodies.</figcaption></figure>

is used to collect biomass directly from Petri plates (see the protocols to extract macromolecules). To this end, sterilized cellophane cut to fit the plate is overlaid onto the medium. Explants are inoculated on it and permitted to grow. On M2, the mycelium will penetrate cellophane after two days of incubation and

**M2 medium.** M2 medium has the following composition: KH₂PO₄ 0.25 g/L, K₂HPO₄ 0.3 g/L, MgSO₄ 0.25 g/L (if MgSO₄ 7 H₂O), dextrin dissolved in warm water 4-6 g/L (this depends on the type of dextrin, we find that potato dextrin at 4g/L works fine, while blond dextrin needs to be added at 5.5 g/L), urea 0.5 g/L, thiamine 0.05 mg/L, biotin 0.05 µg/L and oligo-elements (citric acid 5 mg/L, ZnSO₄ 5 mg/L, CuSO₄ 0,25 mg/L, MnSO₄ 50 µg/L, boric acid (H₃BO₃) 50 µg/L, sodium molybdate (Na₂Mo) 50 µg/L, iron alum (Fe (NH₄)₂(SO₄)₂) 1 mg/L). Adjust to pH=7 with a KH₂PO₄ solution and autoclave for 20 minutes at 120°C.

We find it easier to have stock solutions and to prepare the medium using these stocks. For 1 L: 5 mL of 50 g/L KH₂PO₄, 5 mL of 60 g/L K₂HPO₄, 5 mL of 50 g/L MgSO₄ - 7 H₂O, 5 mL of 100 g/L urea, 0.5 mL of 100 mg/L biotin, 0.5 mL of 100 mg/L thiamine, 1 mL oligo-elements stock solution prepared as follows for 500 mL: 5 g citric acid - 1 H₂O, 5 g ZnSO₄ - 7 H₂O, 1 g Fe (NH₄)₂(SO₄)₂ - 6 H₂O, 250 mg CuSO₄ - 5 H₂O, 50 mg MnSO₄ - 1 H₂O, 50 mg anhydrous H₃BO₃, 50 mg Na₂MoO₄ - 2 H₂O and one drop of chloroform to avoid precipitation of the chemicals. Dextrin is added after dissolving it in hot water.

For Petri plates, agar is added at 10 g/L. Pour 28 ml per Ø=8 cm Petri plate. For race tubes, double the amount of agar (i.e., add 20 g/L). Race tubes are used to measure longevity of the fungus. Designs of race tubes will be explained in the section dealing with Senescence.
will thus be difficult to scrape as early as three days after inoculation, while penetration will not occur for
several days on MR. To collect the mycelium, simply scrape with a sterilized spatula. An additional bonus
of the MR medium is that longevity is much higher and strains seldom undergo senescence.

**MR (Corn Meal) Medium:** MR is produced by dissolving in H₂O 25 g/L of corn flour and 25 g/L of corn cream. Mix and incubate overnight at 58°C (no more and no less). Filter the mix twice with filter papers, then adjust with H₂O again to one liter. Add agar at 12 g/L. Autoclave for 20 minutes at 120°C and pour 28 mL per Ø=8 cm Petri plate.

Other used media are the RG (or M1) medium, an hyperosmotic medium mainly used for protoplasts regeneration (after mutagenesis or transformation), the “Sorbose” medium only used to obtain male gametes (spermatia/microconidia) in high amount and the “Fiole de Roux” medium that is used to produce high quantities of mycelium, especially for protoplast preparation.

**RG medium:** Same as M2 + 205 g/L of saccharose. Autoclave for 20 minutes at 120°C. Pour 28 mL per Ø=8 cm Petri plate.

**Sorbose medium:** Yeast Extract 2 g/L, sorbose 2 g/L, glucose 1 g/L, agar 12 g/L. Autoclave for 20 minutes at 120°C. Pour in tilted assay tubes. Utilization is as follows: inoculate numerous explants per tube and incubate for at least four days. The longer the more spermatia will be recovered. Add 2 ml of sterile water, vortex and filter the suspension with sintered glass filter n°4.

**"Fiole de Roux" medium:** same as M2 + yeast extract 5 g/L. Autoclave for 20 minutes at 120°C. The medium is poured into Roux culture bottles as to obtain about 1 cm at the bottom when laid. It is advisable to add antibiotics in this medium as it is frequently contaminated with bacteria.

Note that dextrin is only one of the food sources that the fungus will eat. Many carbon sources will do. The base is the M0 medium, which is the same as M2, but without dextrin. M3 is M0 + 5.5 g/L of glucose. M4 is M0 + 5.5 g/L of avicel. These can be poured into the Petri plates as the M2 or MR media.
<figure><img src="image_32.png" /><figcaption>**Figure 26** *P. anserina* grows and reproduces on complex carbon sources. The fructifications are the little black dots.</figcaption></figure>

Solid carbon sources, such as hay, miscanthus, wood shavings, paper... can also be used (figure 26). The sterilized food source is added in the plate (usually we use 0.5 g per plate) and 12 mL of sterilized M0 (with or without 12 g/L of agar depending on the need) is poured on it. Strains reluctant to fructify on M2 or M4 may do so on medium with more complex food sources. Hay seems to be particularly handy for this, because many strains obtained from culture collections and that have accumulated mutations, usually yield perithecia on this medium.

In all these media, it is possible to add various metabolites or antibiotics to select for genetically-transformed strains or to allow auxotrophic strain to grow. The concentration for some of them is given in Table 4.
<table><caption>Table 4: supplements to be added to growth media</caption><tr><td>Uridine : 100 µg/mL</td><td>Hygromycin B: 75 µg/mL</td></tr><tr><td>Leucine : 50 µg/mL</td><td>Phleomycin: 10 µg/mL</td></tr><tr><td>Lysine : 100 µg/mL</td><td>Nourseothricin: 40 µg/mL</td></tr><tr><td>Tryptophane : 30 µg/mL</td><td>Geneticin: 500 µg/mL</td></tr><tr><td>Methionine: 30 µg/mL</td><td>Tricyclazol: 1 µg/mL</td></tr></table>

Tricyclazol is used to remove melanin; ascospore without melanin will germinate on all media, but are rather fragile.

As a final note, it is worth stating that *P. anserina* will complete its whole cycle on sterilized horse dung: ascospores will readily germinate on it, mycelium will grow efficiently and fruiting body will be obtained in 7/8 days. However, we have observed some variations with various dung batches; some will permit abundant growth and reproduction, while other will only allow for scarce growth and few perithecia.

## Strains preservation

Long term conservation of *P. anserina* presents no problem. Culture on M2 plates can be kept at 4°C for about two years. To do this, put several explants of the strain to be conserved on a M2 plate. Incubate until the plate is covered with mycelium (if not, contamination by other fungi may occur more frequently). Conserve the stock at 4°C with a parafilm seal. A temperature of no more of 4°C is important, because otherwise residual growth of the fungus will occur and "incolore" mutations will appear at high frequency. These mutations trigger a proliferation of nuclei that carry them to the detriment of wild-type nuclei. Incolore nuclei thus rapidly invade the thalli that become female sterile, lack pigment and produce many spermatia/conidia. For extended conservation, we use cryotubes at -70 to -80°C. To this end, put several explants on a M2 plate. After 2-3 days of growth, take numerous large explants at the growing edge. Store them in a tube containing 1 mL liquid RG (i.e. RG without agar). Place directly the tubes at -80°C. These tubes can be frozen and defrosted numerous times without notable effect on explants regeneration. The strains can be conserved in this way for several years. In my lab, some strains are frozen for nearly 20 years and the explants still regenerate without any problem. Note that some mutant strains may not be stocked as well...
# Methods for macromolecules extraction and genetic transformation

Below are described methods for molecular analyses of the fungus that work well. This list is not exhaustive and many variants for each protocol exist. Note that many of these methods can be used for other species of filamentous fungi. The fungal biomass may conveniently be obtained in two different ways: liquid cultures or Petri plate cultures. For the liquid cultures, shaken Erlenmeyer flasks may be used, especially if aeration is needed. I find that 1 liter Roux flask in which 100 mL of medium is added may be preferable because the flasks can be stacked and shaking is not necessary. Inoculation is made with mycelium fragments obtained with a Fastprep or similar instrument. Calibration of the instrument is required as too much shaking will result in mycelium death... For the Petri plate cultures, we recommend to overlay the medium with a cellophane sheet. The mycelium is inoculated as for the liquid cultures or with mycelium plugs. If the plugs are small, it is not necessary to subsequently remove them. The mycelium will grow on the cellophane and can then be easily scraped with a spatula. However, be aware that the fungus may penetrate the cellophane if incubation is too long (see previous chapter). We usually used liquid cultures when large amount of biomass is needed and Petri plates when small amounts are required.

# DNA Extraction

Depending on the amount and quality of the DNA necessary two different methods may be used: large scale prep or miniprep. The large scale prep will yield large amounts of high quality DNA that can be used for genomic bank constructions (including cosmid banks), sequencing with PacBio or PCR amplification of large DNA regions (> 3 kb). A few samples may be processed in parallel. Miniprep yield is lower and DNA has a lower quality than large scale prep DNA. This DNA does not store as well as the one from large scale prep. It can be used for routine DNA amplification of small DNA fragment (<3 kb), Southern blot analysis or sequencing by illumina. Up to 24 samples may be processed in parallel, although I recommend to
process by batches of 12. Note that a last "instant" method may be used for PCR amplification of small DNA fragment (< 300 bp). It requires minimum manipulation and many samples can be processed in parallel. However, DNA cannot be stored. This last method is used mostly for microsatellite analyses on many samples, as for example required in some mapping analyses.

**For Large scale prep extraction:**

- cultivate the mycelium in Roux flasks in M2 supplemented with 10g/l of Difco yeast extract. Filter with sterile gauze to recover the mycelium and weight it

- freeze in liquid nitrogen. The mycelium can then be kept at – 20°C or – 70°C for extended period

- grind the mycelium with a mortar

- for 2 g of mycelium, add 10 ml of ice-cold TES/Sarkosyl (Tris 12.5 mM, EDTA, 12.5 mM, NaCl, 25 mM pH=8 + Sarkosyl 2.5%) made at the last minute from stocks of Sarkosyl 5% and TES 4x

- incubate for 1h30 at 4°C, mix frequently

- centrifuge for 10 min at 6000 rpm, 4°C

- dialyse supernatant against TES 1x at 4°C overnight

- do alpha-amylase treatment for 2 hours at 20°C (500 µl added from a solution at 2.5 mg/ml in NaCl 3.5M and kept at -20°C)

- do proteinase K treatment for 2 hours at 20°C (50 µl added from a solution at 20 mg/ml and kept at -20°C)

- add 2 vol ethanol 100° and 1/10 vol sodium acetate 3M pH=6

- centrifuge for 5 min at 8000 rpm

- recover in sterile water to make a cesium centrifugation (for a 65VTI rotor, add 4ml sterile water, 4.6 g of cesium chloride and 20 ml of DAPI 1mg/ml, centrifuge for 12h at 45000 rpm). This method allows the separation of nuclear (lower band) and mitochondrial (upper band) DNA.

→ Yield of large scale prep is around 1 mg DNA/Roux flask

**For miniprep**

- inoculate a M2 Petri plate containing a sheet of sterile cellophane. Let grow for 2-3 days

- recover the mycelium with a spatula and put it into a 2 ml eppendorf tube containing 600 µl of TNE/SDS
(Tris 10 mM, EDTA 1 mM, NaCl 100 mM + SDS 2%) to be made from stocks of SDS 10% and TNE 2X

- To break open the mycelium, do 3 cycles of freeze/thaw in liquide nitrogen /water bath at 70°C; vortex between each cycle. Alternatively, if a "Fastprep" or a similar instrument is available, the mycelium may be broken with small glass beads (add ~50 µL of glass beads with Ø= 250-500 µm in the Eppendorf tube). Speed will of course depend upon the used apparatus and must be calibrated beforehand.

- add one volume of phenol pH=8

- centrifuge for 5 min at 4°C 13000 rpm and retrieve the supernatant

- repeat with phenol + chloroform and with only chloroform

- precipitate the supernatant by adding 2 volumes ethanol 100° and 1/10 vol sodium acetate 3M pH=6

- rinse with ethanol 70°, dry and suspend in 40 ml of sterile water

→ Yield of miniprep is a few µg DNA/plate

## For Instant extract

- collect half of a 2 cm thallus grown 48 hours on M2.

- add 20 µL of Ø= 250-500 µm glass beads and 20 µL of freshly-made NaOH 0.5 N

- break open with a calibrated Fastprep

- centrifuge briefly to precipitate the mycelium + beads

- incubate in boiling water for 45 sec – 1 min

- Neutralize with 100 µL of 1:4 Tris 1 M pH= 8.0: TE pH=8.0

→ PCR amplification is done on 1 µL. DNA is very dirty and does not keep more than a few days.

## RNA Extraction

Owing to its fragility, RNA extraction are more difficult than DNA ones. Kits dedicated to the extraction of plant RNA usually work well (such as the RNeasy kits from Qiagen), the key point being to break open the mycelium well enough to recover large amounts of nucleic acids. To do so, it is necessary to ground in a Mikro-Dismembrator (Sartorius, Goettingen, Germany) or similar apparatus for 1 min at 2600 rpm in
vessels containing 9 mm chromium steel grinding balls frozen in liquid nitrogen. Note that apparatuses
with lower grounding strength will give low yields of RNA. For those that prefer using their own devices,
here is a **"hot phenol" extraction** that works well:

- recover the mycelium (from Roux flasks for large scale preps or M2 plates + cellophane for small scale preps)

- freeze in liquid nitrogen

- suspend in 1 or 2 vol of hot phenol made by incubating at 70°C a mix of 1 vol phenol pH=7.5 and 1 vol of tpRNA (NaCl 100 mM, Tris 10 mM pH=7.5, EDTA 1 mM, SDS 0.1%, Sarkosyl 2%), the mix must be homogeneous

- incubate at 70°C for 5 min

- centrifuge for 5 min at 14 000 rpm for small scale preps and 8 000 rpm for large scale preps

- recover the supernatant in a new tube

- repeat by adding phenol + chloroform and then only chloroform

- add 2 vol. of LiCl 6 M and let precipitate overnight at -20°C

- centrifuge for 5 min at 14 000 rpm for small scale preps and 8 000 rpm for large scale preps

→ Yield is low (~50 µg/plate). However, the RNA is very clean and can be used for all usage (northern, cDNA, RT-PCR ...)

**Protein Extraction**

Depending on the intended uses, there are two methods to recover proteins. The first one is intended for Western blotting analyses and the recovered proteins are denatured. The second one is for crude extracts, to measure enzyme activity for example.

**Extraction of denatured proteins for Western blotting**

- recover the mycelium from one M2 Petri plate overlaid with cellophane in a 2 mL Eppendorf tube
- add 50 µL of Ø= 250-500 µm glass beads and 50-200µL of lysing buffer (Tris-HCl pH=7.5 20 mM, NaCl 100 mM, EDTA 1 mM, Triton X100 : 0.05%)

-incubate on ice, while processing all samples

**The following steps must be made very quickly:**

- break open with in a calibrated Fastprep for 40 sec

- add 0.25 vol. of Laemmli charge buffer (Tris pH=6.8 0,4M, 1% β-mercaptoethanol or DTT 50 mM, 1% SDS, 30% Glycerol, bromophenol blue)

- incubate 5 min. at 100°C

- centrifuge 5 min. at max speed in a microcentrifuge

- collect supernatant in a new tube and store at -70°C

-> Use 20 µL per lane for Western blot analysis. Depending on the strain volume should be optimized.

**Crude extract for enzyme assays**

- recover the mycelium from one M2 Petri plate overlaid with cellophane in a 2 mL eppendorf tube

- add 50 µL of Ø= 250-500 µm glass beads and 50-200µL of lysing buffer (Tris-HCl pH=7.5 20 mM, NaCl 100 mM, EDTA 1 mM, Triton X100 : 0.05%)

-incubate on ice, while processing all samples

**The following steps must be made very quickly:**

- break open with in a calibrated Fastprep for 40 sec

- centrifuge 2 min. at max speed in a microcentrifuge at 4°C

- collect supernatant on ice in a new tube and store at -70°C

-> Protein amounts should be assessed before assays are performed.

**Protoplasts preparation**

Protoplasts are mostly used for genetic transformation. They however may also be used in mutagenesis
experiments (see chapter "The sexual cycle and genetical analysis") or to breakdown heterokaryons into
homokaryons. They are prepared according to the following procedure. All steps are performed in sterile
conditions.

- Roux flasks are inoculated with mycelia fragmented with a Fastprep or similar instrument.

- Cultures are grown for 36 h at 27° C in the dark in M2 medium supplemented with 2.5 mg/ml yeast extract ("Fiole de Roux" medium).

- Mycelium is harvested on cheesecloth and washed with TPS1 buffer (0.6 M saccharose, 5 mM Na2HPO4, 45 mM KH2PO4).

- Weight the mycelium. For X g of wet mycelium, add X ml of TPS1 containing 40 mg/ml of Glucanex (Novo Nordisk Ferment AG, glucanex amount may vary upon the efficiency of the batch) and digested for 3 to 4 h at 37° C.

- The protoplasts are separated from mycelial debris by filtration through cheesecloth.

- They are concentrated by 10 min centrifugation at 3200 rpm, washed twice with TPS1 and once with TPC buffer (0.6M saccharose, 10 mM CaCl2, 10 mM Tris, pH= 7.5).

- The final pellet is resuspended in TPC buffer and protoplasts concentration is determined by counting under microscope with a hemocytometer. Protoplasts can be transformed immediately or stored at - 70° C. They can be regenerated in liquid RG medium or on RG plates.

Genetic transformation

- Before transformation, the protoplasts are subjected to a 5 min heat shock at 48° C and transferred 30 secondes on ice and then 5 minutes at room temperature.

- The DNA is added (5 µg of DNA for 0.2 ml of protoplasts at a 10⁸/ml concentration) and protoplasts are incubated for 10-15 min at room temperature.

- Two ml of a PEG solution (60% polyethylene glycol 4000, 10 mM CaCl2, 10 mM Tris pH 7.5) are added and carefully mixed.

- After a 15 min incubation, The protoplasts are centrifuged 10 min at 3200 rpm and then suspended in 5 mL liquid RG and incubated overnight at 27°C.
- The culture is centrifuged 10 min at 3200 rpm and suspended in 500 µL of liquid RG; 100µL aliquots are spread onto M2 selection plates.

Routinely 10 to 50 transformants are obtained per µg of plasmid DNA. Plasmids carrying different selective markers are available for transforming *P. anserina* protoplasts. The *Leu1* or *Ura5* wild-

<figure><img src="image_33.png" /><figcaption>**Figure 27** Plasmids routinely used to transform *P. anserina*. The four plasmids derive from pBluescript pBC-SK+ from Stratagene carrying the chloramphenicol resistance marker. Resistance markers for *P. anserina* were inserted at the unique *Xmn1* site located outside the polylinker. The SK+ polylinker is thus intact allowing for the blue/white selection with the α fragment of the lacZ gene, for the production of single-stranded DNA with the f1 origin, as well as for RNA production with the T3 and T7 promoters. Enzymes cutting once are indicated in blue. The position of the amont and aval primers used for deletion cassette is indicated by the black triangles.</figcaption></figure>
type genes restore growth on minimal medium when introduced into the *leu1-1* and *ura5-6* mutants auxotrophic for leucine or uridine, respectively. Genes allowing selection of transformants resistant to hygromycin B (75-100 µg/mL), phleomycin (5-10 µg/mL), geneticin (250-500 µg/mL) and nourseothricin (40 µg/mL) are also available. These have been placed into the pBC-Hygro, pBC-Phleo, pBC-Genet and pBC-Nourso plasmids to allow convenient cloning of DNA fragments (figure 27).

Note that, while nuclear transformation is easy, introduction of DNA into the mitochondria has never been successfully achieved, despite several attempts made by biolistic as described in *Saccharomyces cerevisiae*.

## Gene deletion, single nucleotide change and at-will modification of the genome

Gene deletion is also fairly straightforward in *P. anserina*. The most convenient way is likely the split cassette method (figure 28). This method starts with two rounds of PCR to construct a two-part deletion cassette. To amplify the selection marker, regions allowing to reproducibly amplify all markers from the pBC vectors were identified (figure 27: amont and aval, Table 5). They can be used to make the fusion primers required to link the flanking regions and resistance markers in the split marker method (the red regions of primers B, C, E and F of figure 28). Noteworthy, they permit to construct deletion cassettes with the four markers using the same set of primers, a feature particularly handy, if one needs the deletion with different markers to construct multiple mutants or combine the deletion with GFP constructs. Once created, the two-part deletion cassette is transformed into *P. anserina*. Inside the fungus, the DNA usually integrates the nuclear genome mostly thanks to the non-homologous end joining pathway. Indeed, only DNA fragments contained in cosmids (hence with a size of about 35 kb) integrate with a frequency of about 80% with the homologous recombination pathway. Fragments even as large as 20 kb integrate with a frequency of 95-99% with the non-homologous pathways. To circumvent this problem, it is advised to use recipients with deletions in the *mus51* and *mus52* genes coding for the Ku70 and Ku80 subunits of the protein complex performing the non-homologous end joining, respectively.

<table><caption>Table 5: amont and aval regions for resistance marker amplification</caption><tr><td>amont</td><td>5'-CTATTTAACGACCCTGCCCTGAACCG-3'</td></tr><tr><td>aval</td><td>5'-CTTACCGCTGTTGAGATCCAGTTCGATG-3'</td></tr></table>
<figure><img src="image_34.png" /><figcaption>**Figure 28** The split marker method for gene deletion. This method is based on two successive rounds of PCRs to construct a two-part deletion cassette, followed by its transformation into *P. anserina*. Three crossing-overs result in the replacement of the gene by the resistance marker. Utilization of recipients deleted for the *mus51* or *mus52* genes coding for the Ku70 and Ku80 subunits that function in the non-homologous end joining of DNA breaks results in a high proportion of *bona fide* replacements.</figcaption></figure>
Deletions for *mus51* and *mus52* are available with all four resistance markers, allowing deletion of any gene with either one of the resistance markers. The *mus51* gene is located on chromosome 6 (CDS number *Pa_5_6180*) and *mus52* on chromosome 7 (CDS number *Pa_7_9250*). Choose carefully which strain to use for deletion of genes on these two chromosomes! After deletion it is strongly recommended to verify the deletion by PCR and Southern blotting analysis, because non-homologous and faulty integration still occurs in the *Δmus51/Δmus52* strains. A good practice is to first select a few potential good candidates by amplifying with PCR the expected function fragments (with the primer pairs “verifamont + X” and “verifaval + Y”). These are crossed with the wild type to clean the strain, *i.e.*, to segregate out the *Δmus51/Δmus52* mutation and any other potential mutations resulting from the transformation procedure. Southern blotting with appropriate probe can then be performed on the F1 progeny.

Interestingly, it is also possible to change a single nucleotide or modify the nuclear genome at

<figure><img src="image_35.png" /><figcaption>**Figure 29** At will modification of the *P. anserina* genome. See text for explanation.</figcaption></figure>
<figure><img src="image_36.png" /></figure>

**Figure 30** Scheme to tag with fluorescent proteins. A 500 bp DNA fragment from the 3'-end of the gene and lacking the stop codon is amplified by PCR with a high fidelity polymerase and cloned into a plasmid containing the CDS of a fluorescent protein, eGFP or mCherry (here the pEGFPhyg1 vector carrying also an hygromycin B resistance marker). Note that pmCherry-Hyg with the same multi-cloning site and resistance marker carrying the mCherry CDS is also available. Integration after a single crossing-over following transformation in a Δmus51/Δmus52 strain results in the expression of the tagged protein from the native site.

will by co-transforming two DNA fragments into the Δ*mus51/Δmus52* strains (figure 29). Indeed, one circular fragment carrying the selection marker allow to select for potential candidates and in a subset of them the linear fragment carrying the desire change integrates by a double crossing-over and changes the sequence as desired. Usually, integration takes place in 5-10% of the transformants, implying that it is better to have a selection scheme for the correct replacement. A genetic cross with the wild type permits then to segregate out the Δ*mus51/Δmus52* mutation as well as the integrated copy of the circular molecule. This method has for example been used to construct PM154, the self-fertile homokaryotic
strain carrying in tandem the two mating type idiomorphs (see chapter “The sexual cycle and genetical analysis”). The final strain differs from the wild type only by the desired modification!

Other genome modifications frequently made are those required for gene overexpression and protein tagging with fluorescent proteins. For overexpression, it is possible to use the constitutive and strong promoters of the *gpd* and *AS4* genes. The *gpd* gene encodes the glyceraldehyde 3-phosphate dehydrogenase, a highly expressed glycolysis enzyme. The *AS4* gene encodes the cytosolic translation elongation factor eEF1A (also known as EF-1α). A region of 300-400 bp upstream of the translation start site from both genes is sufficient to drive high expression. Note that if one wants to express genes from other fungi (for complementation purposes for example), promoters from related *Sordariomycetes* fungi usually work well in *P. anserina*. On the contrary, promoters from the distantly related *Eurotiomycetes* often do not work well. For protein tagging, eGFP and mCherry works fine in *P. anserina* (see figure 34 of the next chapter). Tagging is often made following the strategy depicted in figure 30 that results in the expression of the fusion protein from the native promoter at the endogenous locus. In the wild type, integration occurs randomly in the genome, usually at a single location. However, integration is often

<figure><img src="image_37.png" /><figcaption>**Figure 31** *pCIB2 and pCIB4 plasmids for targeted integration in the genome of P. anserina.* The plasmids carry the nourseothricine resistance marker flanked with the 5' and 3' regions of the *Pa_2_3690* (pCIB2) or *Pa_4_5450* (pCIB4) genes. The *Swal* and *Pacl* sites can be used to clone the cassette/gene to be expressed. The *Ascl* sites can then be used to cut the plasmids. After transformation in a *mus51/52* strain, the linear fragments carrying the 5' region followed by the expression cassette, the nourseothricine resistance marker and the 3' region will replace by a double crossing-over the *Pa_2_3690* or *Pa_4_5450* genes. The *Swal*, *Pacl* and *Ascl* enzymes recognize 8-bp sites with 4 bp overhang to facilitate cloning.</figcaption></figure>
is often preceded by concatenation of several molecule of the transforming DNA, followed by internal
deletions. Correct expression of the transgenes is thus often variable depending on the modalities of the
DNA integration. To ensure a correct expression of transgenes, it is possible to use along with the
*mus51/mus52* strains the pCIB2 and pCIB4 plasmids that target integration at the *Pa_2_3690* and
*Pa_4_5450* loci, respectively (figure 31). This strategy ensures correct expression and even allow for
comparison between transgenes. However, their utilization results in the inactivation of either
*Pa_2_3690* or *Pa_4_5450*. So far, no phenotype has been associated with the inactivation of these genes.

Overall, it is possible to extract macromolecules and modify easily and at will the genome of *P. anserina*. Many additional tools generated by the laboratories working on this fungus are available. Only those routinely used in my lab are presented here.

**Tools for expression analysis**

Although it is still possible to analyze gene expression one gene at a time, tools for transcriptomic analyses have been developed. For example, microarrays of the whole transcriptome are available for strain S. Although, these are based on the first annotation of the *P. anserina* genome, they probe the overwhelming majority of the genes, since few new genes have been discovered since. Transcriptome analyses using these microarrays have been made on *mat+* and *mat*- strains, at various times of ascospores germination, mycelium growth and perithecium development. Most, but not all, of these data are available in public databases. More recently, RNAseq data have been obtained and made public for the following developmental stages: one-day-old and four-day-old mycelia, two-day-old and four-day-old perithecia, non-germinated ascospores and eight-hour-old germinated ascospores. RNAseq data of *P. anserina* grown on two different biomasses are also available. It is likely that additional RNAseq data will be made available in public databases in the near future. All of these data can be searched to have a first idea on the expression pattern of interesting genes.
Morphology and cytology

As stated in the previous chapters, morphological analysis of the sexual reproduction apparatus was the basis for the definition of the species and is still used to differentiate species of the *P. anserina* complex from other fungi. However, it is only a part of the structures that the fungus is able to differentiate during its lifecycle (figure 32). In this section, we will thus examine the morphological features of both the reproductive and vegetative structures elaborated by *P. anserina*.

<figure><img src="image_38.png" /></figure>

**Figure 32** Schematic representations of the major cellular structures differentiated by *P. anserina*. After the germination of ascospore, the fungus differentiates a complex mycelium having different kinds of hyphae. Onto this mycelium, both male and female gametes are differentiated. After fertilization of compatible gametes, the fruiting body or perithecium is formed. In this sexual sporophore, ascospores are produced. They are then expelled to ensure the dispersal of the fungus. The different structures are not to scale.
## The mycelium

The mycelium differentiated by *P. anserina* is typical for a *Pezizomycotina*. Apical hyphae from the outer edge of the thallus (figure 33) have a diameter of about 4-6 µm and contain few nuclei (often 2-5). Very quickly, hyphae branch and anastomose (figure 33). Meanwhile septa are deposited (Figure 34). They contain a central pore associated with Woronin bodies, which are modified peroxisomes involved in regulating the flow within the central pore (figure 34). Once mature, the mycelium is thus a complex network of interconnected hyphae (figure 33). These may have different diameters, suggesting different functions; yet nothing is known about the potential roles of these differently shaped-hyphae. All these hyphae have an internal structure typical for *Dikarya* fungi (figure 34). If in contact with the atmosphere,

<figure><img src="image_39.png" /><figcaption>**Figure 33** Vegetative hyphae of *P. anserina*. Top left, typical apical hyphae; top right; branching pattern of subapical hyphae; bottom left, anastomoses (arrowheads) showing the typical ladder-like morphology; bottom right, typical hyphae from an internal part of the mycelium showing different diameters</figcaption></figure>
<figure><img src="image_40.png" /><figcaption>**Figure 34** Intracellular structures of vegetative hyphae of *P. anserina*. Left, septa with Woronin bodies (arrows); right, some of the structures of the nucleus/reticulum/vesicular network (nuclei in blue, endoplasmic reticulum in green and vacuoles in red. Pictures by S. Brun.</figcaption></figure>

some hyphae will erect, likely thanks to a coating of hydrophobins (figure 35). Additionally, melanin
pigments will accumulate at the center of the colony (figure 35).

At the microscopic level, there is no difference between the mycelia differentiated by the *mat+*, *mat-* and *mat+/mat-* thalli. At the macroscopic level, the presence of fruiting bodies specifically on the *mat+/mat-* thalli appears to modify the behavior of the mycelium with decreased pigmentation and aerial hyphae amounts in the outer edge of the culture (figure 36). Moreover, differences between the *mat+* and *mat-* thalli may be exhibited on some media, especially in interaction with lighting conditions. This is

<figure><img src="image_41.png" /><figcaption>**Figure 35** Aerial hyphae and pigments of 4-day-old *P. anserina* thallus grown on M2. Pigments are partially masked by the aerial hyphae. On M2, these are mostly erected in a ring region located between 1 cm and 2 cm from the center of the culture. This ring region also accumulates more pigment.</figcaption></figure>
<figure><img src="image_42.png" /><figcaption>**Figure 36** Macroscopic morphology of 7-day-old *P. anserina* thallus grown on M2. The *mat+* and *mat*-cultures are older than the one on figure 31 and many aerial hyphae have collapsed onto the medium. The accumulation of pigment is thus more visible, showing the typical ring region. On the *mat+*/*mat*-culture, perithecia are mostly located in the ring region. Note that beyond the ring in the outer edge of the culture, pigment are lacking, while they are present in the inner part of the culture inside the ring.</figcaption></figure>

due to the presence of polymorphic genes on chromosome 1 near the mating type, in a region nearly devoid of recombination. One such gene that has been shown to trigger differences between the *mat+* and *mat*- thalli is *rmp1*. In strain S, the *mat+* thalli carry the *rmp1-2*, while the *mat*- thalli carry the *rmp1-1* allele. Noteworthy, *rmp1-2* confers a slight thermosensitivity. Hence, *mat+* thalli do not produce aerial hyphae when grown at 36°C and grow slightly slower. As seen on figure 31 and 32, there is on M2 a defined ring region with an inner diameter of 1 cm and an outer diameter of 2 cm. It is visible on homokaryotic thalli thanks to melanin and aerial hyphae accumulation and in *mat+*/*mat*- dikaryons thanks to fruiting bodies. How this ring region is produced is still unknown. It is not present on all media and its size and density is agar and carbon-source dependent. Usually the more digestible and abundant the nutrients, the more pronounced this region. For example on the crystalline-cellulose-containing M4, the ring is only visible in *mat+*/*mat*- dikaryons (since *P. anserina* does not produce pigments and aerial hyphae on M4). The ring is wider (inner diameter is 2 cm and outer diameter is 4 cm) and perithecia are much sparser.

When in contact with cellophane, *P. anserina* differentiates special hyphae dedicated to penetrate the solid nutrient layer and digest it from the inside: the appressorium-like structures (figures 32 and 37). These are differentiated after about 12 hours of growth on cellophane and are especially abundant in the ring zone with dense aerial hyphae/perithecia. When inoculated on M0 medium with
<figure><img src="image_43.png" /><figcaption>**Figure 37** Appressorium-like structures of *P. anserina* thallus grown on cellophane. From left to right, different focuses highlight these structures from top to bottom. There is a distance of about 10 µm, between each picture. First on the left, hyphae grow parallel to cellophane. Then hyphae reorient and differentiate the appressorium-like structures (black arrowhead) that contact the solid layer of cellophane. From these, hyphae with a tiny diameter called penetration pegs (white arrows) penetrate the cellophane. Then on the right, haustorium-like structures develop inside the cellophane. From these, new penetration pegs (white arrow) descend further inside the cellophane layer.</figcaption></figure>

cellophane, the fungus will cross the cellophane layer (about 50-100µm) in about three days. These structures are inhibited by the presence of glucose in the medium. It is presently not known if they are also differentiated when *P. anserina* grows on natural biomass, especially when growing on dung.

In addition, to the hyphal structures described above, the friendly mold is able to differentiate
lasso-like hyphae, whose role is unknown (figure 38). The fungus also differentiates in media poor in

<figure><img src="image_44.png" /><figcaption>**Figure 38** Some vegetative structures of unknown function differentiated by *P. anserina* thalli. Left, lasso-like hyphae; right, miscrosclerotic structures.</figcaption></figure>
nutrients, such as M4 and wood shavings, hay or miscanthus media, small sclerotic-like structures having a diameter of about 50 µm (figure 38). The exact nature as well as the role of these structures is also unknown. *P. anserina* may also be able to form sclerotes with larger sizes, however, these are seldom observed and conditions enabling their production have yet to be defined.

The gametes

*P. anserina* differentiates both male and female gametes on the mycelium and hence display an “ascohymenial” development. On M2, differentiation starts after two days of growth and fertilization can proceed on 3-day-old thalli, showing that both types of gamete are functional at that time. Both the *mat+* and *mat-* thalli differentiate the two kinds of gametes. In fact, we know that the mating type locus is

<figure><img src="image_45.png" /><figcaption>**Figure 39** Differentiation of the *P. anserina* female gametangium. The fertile tissue is colored in a lighter color compared to the maternal one. See text for description.</figcaption></figure>
<figure><img src="image_46.png" /><figcaption>**Figure 40** Differentiation of the *P. anserina* male gametes. Usually, spermatia are born at the apex of a peg developing from a spermogonium. Repetitive budding produces small cluster of spermatia. Under some conditions (here top middle and top right in the dark at 18°C on M2 + cellophane), a proliferation of spermogonia and spermatia occurs. Erected sterile hyphae also differentiate in these structures that strikingly resemble *Cladorrhinum/Bahupaathra* anamorphs. P: protoperithecia, arrow shows a cluster of spermatia/conidia in the anamorphic-like structures. See text for details.</figcaption></figure>

dispensable for gamete differentiation.

Female gametes - or more properly gametangia since they are pluricellular when mature - are typical ascogonia/protoperithecia (figure 39). Differentiation starts with characteristic hooked hyphae that will continue their differentiation by generating a pluricellular and multinucleated ascogonium coils. These coils will then be embedded by neighboring hyphae to generate protoperithecia that may reach large sizes (some exceed 100 µm). At that stage, the gametangia are already composed of two different tissues: the female sexual tissue awaiting fertilization in the center originating from the ascogonium coil and that reaches the outside thanks to special hyphae called trichogynes, and several layers of sterile maternal tissue that protects the sexual tissue and that will shape the fruiting body. This sterile tissue has
thus a maternal origin and results from the proliferation of the hyphae that surround the ascogonium coil. In *P. anserina*, the protoperithecia (and later perithecia) have protruding hyphae that may be mistaken for trichogynes. It is therefore not known whether a single or multiple trichogynes is/are produced per fruiting body.

Male gametes are differentiated on pegs emerging from special hyphae called spermogonia (sing. spermogonium, figure 40). They are small spherical cells having a diameter of 2 µm. They contain a single nucleus and a single mitochondrion. They are produced in small clusters and are very easily released from the spermogonium so that it is very difficult to see them attached to the peg (see figure 40 top left). Attempts to germinate spermatia have so far failed. Indeed, although a paper reported their germination at a rate of 1% on media containing yeast extract and sorbose, this rate was not reproduced recently on this medium and germination proceeds at a frequency of 1 out of one million spermatia in all tested conditions. Intriguingly, in some situations, *P. anserina* differentiates *Cladorrhinum/Bahupaathra* anamorph structures (figure 40, see figure 11 for comparison). Here, there are dozens of "spermatia" carried on "spermogonia", questioning whether these are in fact "conidia" on "phialides" (conidiophores with peg-like structures to carry the spores). Indeed, their sheer number suggests that they may be used for dispersal rather than fertilization. However, these also germinate at a very low rate. Until conditions that reproducibly trigger their germination are found, these are thus better seen as spermatia.

Once the gametes are differentiated, fertilization occurs rapidly between mating compatible gametes (see chapter on the sexual cycle and genetical analysis). Based on experiments with successive waves of fertilization with genetically-different nuclei, fertilization is rapid and efficient since most female gametes are fertilized in 10 minutes. During this event, the nucleus from the spermatium enters the trichogyne and migrates to the ascogonium body. This is followed by the rapid development of the perithecium.

## The perithecium

Maturation of the perithecium from fertilization to the first expulsion of ascospores lasts about four days on M2 medium at 27°C. While unfertilized protoperithecia may reach large sizes, typically about 50 µm, sometime more, they never present the neck and ostiole characteristic of fertilized perithecia and through which ascospores are expelled (see figure 41). As stated above, protoperithecia are composed of
<figure><img src="image_47.png" /><figcaption>**Figure 41** Differentiation of the *P. anserina* perithecium. Top, overall morphological evolution during perithecium development with timeline of main events. Bottom, schematic representation of sectioned perithecia with main tissues.</figcaption></figure>

three to five layers of flattened and pigmented cells protecting the ascogonium that is embedded with sterile maternal tissues. Hairs and trichogynes emerge from the protective layer: at that time the protoperithecium looks thus like a small urchin (figure 41). Upon fertilization, development proceeds and the perithecium enlarges (figure 41). The protective cells differentiate a peridium having two layers: in the inside, the endostratum formed by hyaline polygonal cells of about 12-25 µm, and, on the outside, the exostratum formed by melanized thick-walled polygonal cells of about 10-20 µm. Twenty four hours after fertilization the neck initial is visible on the top of the developing fruiting body as a small area with
increased pigmentation. Meanwhile, the first paraphyses differentiate as early as 24h after fertilization. They are elongated unbranched hyphae composed of several articles. They occupy the centrum along with the first binucleated ascogenous hyphae emerging from the fertilized ascogonium. These produce croziers about 30 hours after fertilization and the first meioses occur just after (figure 37). Two days (48 h) after fertilization the centrum is a mix of croziers, young asci at different stages of development and upward paraphyses (figure 41 and 42). The largest increase in size occurs at day 3 and day 4, at which time the perithecium reach a diameter of about 300-350 µm. The neck is growing upward and the ostiole is forming. On the outside, the cells of the neck undergo intense pigmentation and a small tuff of darkly pigmented hairs (the setae) appears at the end of the third day on many but not all fruiting bodies. On the inside, the neck is lined with special cells, the periphyses. The centrum proliferates, producing more and more asci. These mature and the first delimitated ascospores are seen 54 hours after fertilization. Rapidly, they enlarge, mature and get pigmented. The centrum occupies then the largest part of the fruiting body and is easily separated from the peridium.

The development of the sexual tissues is typical for the *Pezizomycotina* (figure 42). The fertilized ascogonium produces several multinucleated cells from which binucleated cells emerge: the ascogenous hyphae. These binucleated cells have two genetically different nuclei and are thus dikaryotic (see chapter on the sexual cycle and genetical analysis). The dikaryotic binucleated state is propagated during cell divisions thanks to the special divisions of the croziers. The apex of this cell forms a hook. After the

<figure><img src="image_48.png" /><figcaption>**Figure 42** Development of the *P. anserina* sexual tissues. See text for explanation. Right, picture V.</figcaption></figure>

Berteaux-Lecellier.
<figure><img src="image_49.png" /><figcaption>**Figure 43** Mature *P. anserina* fruiting bodies.</figcaption></figure>

coordinated division of the two nuclei, two
septa form and individualize a binucleated
dikaryotic cell that becomes the ascus and
two uninucleated cells that undergo cell
fusion to restore a binucleated dikaryotic cell.
This cell will divide further to produce new
croziers. Meanwhile, the ascus enlarges,
undergoes karyogamy, then immediately
meiosis and a postmeiotic mitosis. The
orientation of the division spindles is parallel
to the axis of the ascus during the meiotic
division and transverse in the postmeiotic
mitosis. Then nuclei glide as to associate two
by two non-sister nuclei. The process ends by
the differentiation of the ascospores that will
be the subject of the next section. Once

mature (figure 43), perithecia may produce a few hundreds to a thousands of ascospores for three to ten
days depending upon the growth medium and humidity of the atmosphere.

The asci and the ascospores

Ascus and ascospore maturation lasts
for about two to three days on M2 at 27°C.
After bursting open mature perithecia and
collecting the centrum, the whole sequence of
asci and ascospore maturation may be seen in
the rosettes (figure 44). Just after their
delimitation around two nuclei by two
membranes, ascospores have a club shape

<figure><img src="image_50.png" /><figcaption>**Figure 44** *P. anserina* rosettes of asci with paraphyses.</figcaption></figure>
(figure 42 & 45). Rapidly, the two nuclei undergo mitoses to yield four nuclei, three of which remain in the head of the ascospore and the fourth one migrates into the tail (figure 45). A septum is then laid down between the head and tail parts of the ascospore when melanin is deposited in the head cell. The nuclei of the tail cell degenerates and the tail cell dies. This dead cell becomes the primary appendage of the ascospore that gives the name to the genus *Podospora* (spore with foot). Meanwhile, the three nuclei of the head part undergo further mitoses and the cell enlarges. When mature, ascospores have around thirty nuclei. The cell wall of the head cell gets heavily melanized by the deposit of three successive layers of melanin, giving it a black color. At least, two gelatinous appendages, the secondary appendages, are laid at the opposite poles of the spores: one at the distal tip of the tail cell and one near the apex of the spore head. They consist of gelatinous materials that are deposited between the two membranes of the

<figure><img src="image_51.png" /><figcaption>**Figure 45** Development of the *P. anserina* ascospores. See text for explanation.</figcaption></figure>
spores. Two smaller ones may also be present on the primary appendage. Their role is unknown. In addition, the four ascospores in the ascus are linked together by a rope of actin that is the remnants of the actin fibers that allow migration of nuclei during the maturation of the ascus and scaffold the delimitation of the ascospores. The last feature of the ascospores is the presence of a germ pore at the apex through which germination will proceed. When mature, the spore head is ~19 x 35 µm and the primary appendage (also called the pedicel) is ~5 x 20 µm.

Asci measure about 25 x 200 µm. Their apices have a simple structure (figure 46). They typically contain four dikaryotic or “big” ascospores (figure 44 & 46). However, asci with abnormally-delimited ascospores are frequent. Their frequency can be increased by incubating developing perithecia with anesthetics such as chloroform (to do so, a cotton wool dipped in the anesthetics is placed along with the Petri plate carrying the developing perithecia in a sealed container). They are due to defects in the positioning of the spindles during the meiotic and postmeiotic divisions, resulting in abnormally-placed nuclei. Indeed, ascospore delimitation is driven by the spindle pole body. If far apart, nuclei will be incorporated in different ascospores, while close nuclei are packed inside the same ascospore. The most common is the five-spored ascus, with three big dikaryotic ascospores and two “small” homokaryotic ones (figure 46). Often, the two small ascospores stem from the alteration of a big ascospore. They result from defects in a single postmeiotic spindle resulting in two sister nuclei being too far apart. However, more complex situations may be found, such as the one depicted in figure 46, where the two small

<figure><img src="image_52.png" /><figcaption>**Figure 46** *P. anserina* asci. Left, ascus apex. Right, arrow points towards a five-spored ascus.</figcaption></figure>
<figure><img src="image_53.png" /><figcaption>**Figure 47** *P. anserina* asci after expulsion from the perithecium. These two asci were collected onto an agar plate laid over a cross plate as depicted on figure 23.</figcaption></figure>

ascospores and the big ascospore between them arise from the bad delimitation of two postmeiotic spindles. Two centrally-placed nuclei formed the big ascospore and the left over nuclei the small ones. Asci with two big and four small ascospores, as well as one big and six small or even eight small ascospores are also encountered, the first one being fairly common, while the last type with eight ascospores being extremely rare. Other atypical asci carry "giant" ascospores resulting from the formation of the ascospores around three, four, five, six, seven or even eight nuclei, the more nuclei the larger the ascospore. All combinations of giant, normal and small ascospores may be encountered. However, they are rather rare. For example, delimitation encompassing eight nuclei yields ascospore with the shape of a banana. They are seen only if anesthetics are applied. The most frequently atypical asci with giant ascospores encountered in usual conditions is two big, one small and one giant ascospore. Frequency of abnormal asci depends upon many factors including the medium, the temperature and the strain. For example our reference strain "S", like most *P. anserina* strain, has a frequency of 5-spored asci of about 1% and giant ascospore-containing asci of about 1‰. Usually, abnormal asci are more abundant early in the production of the ascospores by the perithecium.

Once mature, the four ascospores of an ascus are expelled together, because they are linked with an actin rope as stated earlier (figure 47). A film depicting this event can be seen at the following URL: https://doi.org/10.1371/journal.pone.0003237.s002. Average expulsion speed is 21 m/s, enabling to eject the ascospores about 20 cm away. To do so, ascospores are accelerated with the staggering rate of
<figure><img src="image_54.png" /><figcaption>**Figure 48** *P. anserina* ascospore germination. See text for details.</figcaption></figure>

1 500 000 m/s²! Surprisingly, this does not require extravagant turgor pressure. Indeed, presence of erythritol and pinitol at a combined concentration of 70 mM appears sufficient to do the trick and create the required pressure of 0.4 MPa.

After their landing, ascospores await a trigger for their germination (see chapter Physiological and molecular analysis: deciphering developmental pathways - Ascospore germination). They usually do so by extruding a germination vesicle (also called peg) from the germ pore a few hours after being triggered to germinate (figure 48). From this vesicle, hyphae rapidly emerge to create a new mycelium. Movie n°3 shows these events in an accelerated manner. Note that damaged ascospores may germinate from the breakage points. In addition, few immature ascospores are sometime ejected from the perithecia; these may also germinate from any part of the ascospore.
# The sexual cycle and genetical analysis

## The lifecycle of P. anserina

In the previous chapter, we have studied the various structures differentiated by *P. anserina*, especially those involved in the sexual reproduction. We shall now see how these differentiations are orchestrated during the life cycle of the fungus. *P. anserina* is a heterothallic “pseudo-homothallic” filamentous ascomycete, whose haplobiontic sexual cycle displays interesting features (figure 49). The fungus has thus mating types, i.e., a system of genetic incompatibility/compatibility for fertilization, yet

<figure><img src="image_55.png" /><figcaption>**Figure 49** *P. anserina* life cycle. See text for details. I thank D. Zickler for the original drawing of this cycle.</figcaption></figure>
produces ascospores generating mycelia able to mate with themselves.

Under appropriate conditions (see the section on Physiological and molecular analysis: deciphering developmental pathways – Sexual reproduction), mycelia of each mating-type (*mat+* and *mat-*) are able to differentiate both the male (spermatia) and the female (ascogonia) "organs". However, fertilization can only occur between a spermatium and an ascogonium of opposite mating-types. It works in both directions: ♂ *mat-* x ♀ *mat+* and ♂ *mat+* x ♀ *mat-*. To do so, the ascogonium produces a trichogyne, a dedicated hyphae that is attracted to the spermatia thanks to a pheromone/receptor system and "catches" them. The pheromone produced by the spermatium is specific for each mating type as is the compatible receptor expressed in the trichogyne. Once a spermatium caught, its fertilizing male nucleus enters the ascogonium. A trichogyne is fertilized by a single male nucleus, because to date ascogonia fertilized by genetically different nuclei have never been seen. Note that nuclei from the vegetative mycelium can also be used as male gametes, because fragmented mycelia lacking spermatia are able to fertilize compatible ascogonia. As stated in the previous chapter, an interesting point is that this fertilization event is not immediately followed by nuclear fusion or karyogamy. The two nuclei first divide in a common cytoplasm to form a syncitium. The resulting nuclei migrate by pairs of opposite mating type in specialized cells: the ascogenous hyphae. These hyphae give rise, after divisions, to the crozier cells, where the two nuclei divide synchronously. A special septum is formed to yield three cells. Karyogamy takes place in the upper binucleated cell. It is immediately followed by meiosis, a postmeiotic mitosis and formation of asci. Therefore, many asci are produced after a single fertilization event. Note that all events following a single fertilization take place in a single perithecium, because a single fertilization has never been reported to produce more than one fruiting body.

As stated in the previous chapter, asci usually contain four big ascospores or, in 1% of the cases, five, i.e., three big and two small ascospores. The big ascospores usually carry *mat+* and *mat-* nuclei and thus generate mycelia able to self-fertilize, hence the term pseudo-homothalism used to characterize the breeding system of *P. anserina*. Small ascospores, but also few large ascospores, carry nuclei of a single mating type. They produce homokaryotic self-sterile mycelia that will need a compatible partner of opposite mating type to carry out sexual reproduction. Such large ascospore may be present in five-spored asci resulting from defect in more than one spindle (as the one in figure 46), but also in four-spored asci in which the mating type locus has undergone a first division segregation (see below). Availability of both heterokaryotic and homokaryotic mycelia entails that there are several ways to set up
<figure><img src="image_56.png" /><figcaption>**Figure 50** Three frequent setups for *P. anserina* crosses. See text for details.</figcaption></figure>

sexual crosses with *P. anserina* (figure 50). Self-fertile mycelium (obtained from a *mat+/mat-* big spore or by mixing *ma+* and *mat-* fragmented mycelia) may be inoculated at one or more locations in a Petri plate. This will give rise to the typical ring of perithecia mentioned at the end of the chapter "Podospora anserina" in the tree of life: classification of the species" (see figures 19 and 50). Alternatively, homokaryotic mycelia can be crossed with various setups. The two major ones are "confrontation" and "confrontation arrosée". In the first one, plugs for compatible mycelia are deposited 2 cm apart and incubated without further manipulation. This will result in the production of a line of perithecia at the confrontation between the perithecia (figure 50). Confrontations are for example used to test for the mating type (figure 51). In "confrontation arrosée", three days after setting the plugs, some sterile water (2 mL for an 8-cm Petri plate) is added and the plate is gently shaken. This will dislodge spermatia but not ascogonia. Fertilization will thus specifically proceed with ♀ *mat+* × ♂ *mat-* on one side of the plate and ♂
<figure><img src="image_57.png" /><figcaption>**Figure 51** Typical mating type tests. On this plate, *mat+* and *mat*- tester strains (in white) were confronted to eight thalli to ascertain their mating type. Deduced mating types are indicated in red.</figcaption></figure>

*mat+* x ♀ *mat*- on the other side (figure 50). In *P. anserina*, it thus very easy to orient the crosses as desired, in order for example to study cytoplasmic versus nuclear segregations. To do so, one needs only to do "confrontation arrosée" and remove with a scalpel the mycelium used as male. Ascospores can then be recovered as usual from the remaining female thallus.

An important feature of *P. anserina* sexual cycle is its rapidity. Overall, it takes about eight days to complete the cycle from ascospore germination to the next generation of ascospores. Ascospore germination occurs overnight (on G medium or after a heat shock). If inoculated on M2, three days after gametes are differentiated. Finally, the process from

fertilization to ascospore maturation takes in optimal condition about 4 days at the end of which the ascospores are ejected out of the perithecium. This is much faster than *Neurospora crassa* and *Aspergillus nidulans*. In these two species, the complete cycle lasts two to three weeks. Ascospore production by *P. anserina* crosses lasts for about three to ten days depending upon the medium used and humidity of the incubation chamber. Plates producing ascospores may be stored at 4°C, but no more than a few days (usually at most three days), because ascospore ejection won't resume if longer incubation time is used. Ascospores stored on "Agar plates" will germinate with 100 % efficiency when collected rapidly. However, germination efficiency will drop after three to four weeks at 4°C. Similarly, mycelia issued from germinated ascospores won't keep more than a few weeks on G medium at 4°C, while they may last two to three years on M2 medium at 4°C.

Sexual reproduction appears to be the principal mean of dispersion of *P. anserina* as it seemingly lacks any special mean of asexual reproduction, especially the fertilizing spermatia do not seem to germinate (see previous chapter). However, fragmentation of the mycelium will regenerate new thalli able to complete the whole lifecycle. This is a routine procedure used in the labs. Note that although it is frequently assumed that any cell from fungi is totipotent and could regenerate thalli fully able to
complete the whole lifecycle, spermatia, but also ascogonia and also many cells from the centrum of the
perithecia appear unable to do so in *P. anserina*. This has been reported by some of the older authors
working on *P. anserina* development but also observed by Robert Debuchy and myself.

Mutant generation

UV mutagenesis at 254 nm is the most convenient method to recover mutants. Doses for wild
type range from 100 J/m² to 300 J/m², the best dose being usually 200-250 J/m². It corresponds to 1%
survival when measured with protoplasts regenerative potential. Screens obviously depend upon the
desired mutants. Positive screens may be started from fragmented mycelium. UVs are usually applied
one day after inoculation when the mycelium fragments have started to regenerate and form a "veil" of
very young mycelium on the plate. However, if no mutants are obtained with a direct screen or a
negative screen is to be used, there is still the possibility to do mutagenesis on protoplasts. Most are
uninucleated (and haploid), hence any mutation will immediately express its phenotype. To do this, one
needs to do protoplasts as indicated in the section "Methods for macromolecules extraction and genetic
transformation - Protoplasts preparation"; then one spreads them onto RG medium at a concentration of
10⁴ per plate. Irradiate with UV at 200 J/m². Incubate in the dark at 27°C. Regenerants can be picked up
after 48h under the binocular to test them individually. At least 5000 of them should be scrutinized
before giving up!

Another way to recover mutants is to tape into the genetic diversity of the *P. anserina* species
complex. Indeed, some species may present interesting features not present in *P. anserina*, such as the
development of "sectors", spore killer genes, the ability to use some carbon sources more efficiently, etc.
It is possible to analyze the genetic bases of these differences by crossing the interesting species with the
sequenced reference strain S (or s) of *P. anserina*. Fertility of these interspecific crosses is poor, but it is
possible to obtain few ascospores that in turn will have a better fertility when crosses with both parental
strains. It is thus possible to start an "isogenization" process during which the interesting genetic
difference (hopefully due to a single polymorphism) can be progressively associated with more and more
of the genome of the *P. anserina* reference strain S. We usually do 10 backcrosses with strain S, at which
time, the final isogenized strain contains the polymorphic locus and few neighboring genes from the
other species, while the rest of its genome comes from S.
## Genetical analyses: segregation, dominance/recessivity and complementation

As stated in the previous chapter, spindle positioning and ascospore delimitation is completely defined in *P. anserina* asci and can be summarized as depicted in figure 52. This entails some interesting features for genetic analyses. Firstly, if no crossovers occurred between a gene (with a+ and a- alleles involved in the cross) and its centromere, the ascus contains two ascospores with two a- nuclei and two with two a+ nuclei (see how centromeres behave in figure 52). Secondly, if a crossover has occurred between the gene and its centromere, the ascus contains four ascospores with one a- nucleus and one a+ nucleus (see how the mating type idiomorphs behave in figure 52). Therefore, first (FDS) and second (SDS) division segregation frequencies can be easily calculated (figure 53). Note that this allows doing ordered tetrad analysis with disordered tetrad!

<figure><img src="image_58.png" /><figcaption>**Figure 52** Meiosis and gene segregation in *P. anserina*. Centromeres are the black and brown dots. Only chromosome 1 carrying the mating type locus is represented.</figcaption></figure>
<figure><img src="image_59.png" /><figcaption>**Figure 53** First division (FDS) and second division (SDS) segregation in *P. anserina*.</figcaption></figure>

A second important feature of *P. anserina* meiosis is the positive interference exhibited by crossing-overs, i.e., the presence of one crossing-over inhibits the occurrence of a second one nearby. The interference is so powerful in *P. anserina* that only one crossing-over usually happens over large regions. Especially, in 98-99 % of the meiosis, a single crossing-over occurs between the mating type and its centromere on chromosome 1, allowing for the recovery of *mat+/mat-* ascospores yielding self-fertile mycelia in nearly all asci. Note that on this chromosome arm a second crossing-over usually takes place between the mating type locus and the telomere. This situation is encountered for many chromosomes, with a first crossing-over between the centromere and a region located at about two thirds of the chromosome arms and a second one more distal. Therefore, SDS frequencies usually increase as the genes are positioned farther away from the centromere then decrease to reach the 66 % usually seen
<figure><img src="image_60.png" /></figure>

**Figure 54** Linkage map of *P. anserina*. Positive interference of crossing-overs results in most genes grouped together around the centromere and in the middle of the chromosome arms. The black dots are centromeres. Further analyses carried out since this last official version of the map was published in 1982 have filled some of the gaps between the clustered genes (see figure 63 for an updated version of the genome map). The “left” arms are at the top and the “right” ones at the bottom.

when multiple crossing-overs are present other the same chromosome arm. This entails a linkage map
quite different from the ones of other model organisms (figure 54) and the possibility to easily order
genes along the chromosome arms. Indeed, parental ditypes, tetraypes and non-parental ditypes are
easily determined (figure 55). When two genes are on the same chromosome, non-parental ditype are
rare. Moreover, if they are on the same arm, one of the tetratype asci will be more frequent: the one
with the gene closest to the centromere segregating with FDS and the most distal with SDS (figure 55).

*P. anserina* having a haplobiontic life cycle, vegetative diploids are difficult to select and are highly unstable (see the next section on the parasexual cycle). So, apart from the mutations that act between fertilization and meiosis, dominance/recessivity tests must be performed with heterokaryons. There are several simple ways to obtain heterokaryons. They can be directly obtained after sexual reproduction from the big spores. Mycelia of both mating type may be mixed together by simply grinding them in an Eppendorf tube with a spatula or by using a shaking apparatus such as a Fastprep and then
<figure><img src="image_61.png" /><figcaption>**Figure 55** Segregation of two couples of alleles in *P. anserina*. PD: parental ditype, NPD: non-parental ditype and T: tetratype.</figcaption></figure>

inoculating the mix on a fresh Petri plate. Note that these tests in heterokaryons can produce some
interpretation problems since very often mutant nuclei (sometime wild-type ones) can be easily lost. To
circumvent this, there are simple genetics tricks. One can use strains with complementary mating types,
allowing checking through the presence of perithecia if both nuclear types are present. Association of the
mutation to be tested with the *leu1-1* (or *lys2-1*) mutations will permit to make forced balanced
heterokaryons with *lys2-1* (or *leu1-1*). Testing on minimal medium will ensure that both types of nuclei
are present. Note that a problem frequently encountered in heterokaryon construction is the "vegetative
incompatibility". Indeed, when anastomoses occur between thalli of the same strain, heterokaryons will
readily form and proliferate. However, when anastomoses occur between different wild-type strains,
frequently heterokaryotic hyphae will die in a violent death reaction called incompatibility reaction (see
section on "Hyphal Interference and Vegetative/heterokaryon Incompatibility"). This occurs for example
**vegetative diploids**

<figure><img src="image_62.png" /><figcaption>**Figure 56** Vegetative diploids using the SP40 mutant. Taken from the thesis of V. Berteaux-Lecellier</figcaption></figure>

when the S (BIG S) strain is confronted with the s (small s) strain. These two strains are the most commonly used for genetical analysis! In summary, it is advised to construct heterokaryons with compatible strains! Note that complementation tests are often carried out with heterokaryons as described above for dominance/recessivity.

## Parasexual cycle

P. anserina, like many ascomycete fungi, is able to form vegetative diploids and hence undergoes a parasexual cycle. However, as stated above, vegetative diploids are quite difficult to obtain. Indeed, they are rare and unstable. There is a way to visualize them by using the SP40 mutant that carries a reciprocal translocation. When it is crossed with the wild type, some big ascospore progeny gives rise to spindly unpigmented thalli. In such thalli, the two nuclear components carry unbalanced sets of chromosomes. Hence, homokaryons would be non-viable and both types of nuclei must stay in the thalli for their survival. However, in the spindly thalli, complementation between the nuclei is weak, resulting in poor growth. On these spindly thalli, dense patches of pigmented mycelium frequently appear (figure 56). These result from vegetative karyogamy followed by "haploïdization" of the nuclei. This occurs *via* chromosome loss restoring a balanced set of seven nuclear chromosomes. Mitotic crossing-overs as described in the parasexual cycles of other fungi may also occur. Note that the sexual cycle of P. anserina is so efficient that the parasexual cycle is used only in special conditions, such as to test whether karyogamy can proceed in some mutants. Moreover, the SP40 strain seems to have been lost from collection...

## Developmental genetics: grafting and genetic mosaics
<figure><img src="image_63.png" /></figure>

**Figure 57** Grafting of perithecia to study their development in *P. anserina*. Nine 2-day-old wild-type perithecia were placed onto a homokaryotic wild-type thallus (left). Two days later (right), transplanted perithecia have completed their development.

Because fruiting body development is the most complex differentiation process exhibited by *P. anserina*, several tools enabling to analyze it genetically have been designed. Firstly, developing perithecia may be transplanted onto other genetically-identical or genetically-different mycelia (figure 57). To do so, simply lift a perithecium along with the smallest amount of underneath agar and gently put it onto the desired mycelium. This should be optimized as taking too little underlying agar will result in perithecium drying and bursting, while taking too much may obscure the results (see movie n°1). When wild-type perithecia are transplanted onto wild-type mycelia, development resume as if nothing happened and nearly 100 % of the grafted fruiting bodies will mature in four days as control non-grafted ones. Note that success rate with one-day-old perithecia may be lower since at that stage it is still difficult to differentiate fertilized from unfertilized protoperithecia. As seen in figure 58, if transplanted onto mutant mycelia, the fruiting body may or may not complete development depending whether the gene affected in the mutant is required or not in the mycelium. The same reasoning applies for mutant perithecia transplanted onto the wild type: maturation will proceed only if the gene affected in the mutant is dispensable in the fruiting body. Note that this kind of experiments with perithecia from mutants is not frequently done because some fruiting body development is needed for grafting to proceed!

Mosaic analyses with the *Δmat* and *pks1-193* mutants enables to gain further insight into the role of genes during development. The *Δmat* strain is a mutant for which the mating type locus has been
<figure><img src="image_64.png" /><figcaption>**Figure 58** Grafting of perithecia to study their development in *P. anserina*. See text for details.</figcaption></figure>

deleted. This strain is unable to engage fertilization, yet its mycelium is like that of the wild-type. It can thus be used to complement mycelium defect(s) impairing perithecia development. Noteworthy, it is able also to complement maternal defects, but of course not zygotic ones. Hence, when *mat+/mat-* heterokaryotic mutant mycelia are mixed together with *Δmat* in a trikaryon, development of perithecia on the trikaryon means that the gene affected in the mutant is dispensable in the zygotic lineage, while lack of fruiting body entails that this gene is required in the zygotic lineage (figure 59). Note that analyses of the defects in the trikaryons may also enable to dissociate maternal and zygotic defects for genes acting at multiple steps during development. Mosaic analyses with *pks1-193* permit to address whether the development defects are due to mycelium or maternal tissues defects. The *pks1-193* mutants carry a mutation in the gene encoding the polyketide synthase that acts at the first step of the biosynthesis of
<figure><img src="image_65.png" /><figcaption>**Figure 59** Trikaryon analyses with Δ*mat* to study development in *P. anserina*. see text for detail.</figcaption></figure>

melanin. In *P. anserina* this mutation is cell autonomous, because melanin pigments do not diffuse in
hyphae. Also the gene acts at all stages of *P. anserina* development: ascospores, mycelia and fruiting
bodies, all lack pigments in the mutant (fruiting bodies are also slightly smaller in the mutant). When
heterokaryons of compatible mating types are made between the wild type and *pks1-193*, two kinds of
perithecia may be observed: pigmented (or dark) ones and non-pigmented (or light) ones (figure 60). The
maternal tissues of the pigmented perithecia stems from the wild type and from *pks1-193* in the non-
pigmented ones. Note that very few mosaic perithecia with part of their peridium pigmented and part
non-pigmented are observed. This suggests that few nuclei are involved in setting up the differentiation
of the maternal tissues of the perithecia. When the mosaics are made with a developmental mutant
lacking the ability to differentiate fruiting bodies (i.e., a mutant x *pks1-193* heterokaryon of compatible
<figure><img src="image_66.png" /><figcaption>**Figure 60** Mosaic analyses with *pks1-193* to study development in *P. anserina*. Arrows: dark (=pigmented) perithecia. See text for detail.</figcaption></figure>

mating types), two different outcomes may usually be observed (figure 60). Firstly, no pigmented
perithecia may be recovered. This suggests that only perithecia with *pks1-193* maternal tissue may
develop and hence that the gene affected in the developmental mutant is required in the developing
perithecium. On the contrary, if both pigmented and non-pigmented perithecia are observed, this
indicates that the altered gene is not required in the developing fruiting body. Note that a third situation
may be observed: the presence in high number of mosaic perithecia with part of their peridium
pigmented and part non-pigmented. This is accounted for if the product encoded by the mutant gene can diffuse from cell to cell and hence in this case the wild type allele of the gene is not required in all the cells of the fruiting body.

The last tool generated to study development is the *PM154* strain. This strain carries both mating type idiomorphs (*mat+*_mat-) in tandem at the mating-type locus on chromosome 1, without any additional genetic modification (figure 61). It is self-fertile and able to fertilize both *mat+* and *mat*-strains. Although it generates abnormal progeny with many incomplete asci, it can be used to easily select sterile mutants, including those affected with recessive mutations in genes acting in the zygotic tissues. Note that in crosses

with *mat+* and *mat-* strains, there is a large distortion of segregation with the preferential recovery of *mat+*_mat-* progeny.

## Genes and genome

With the advent of sequencing and genetic transformation, genetic analyses in many organisms including *P. anserina* have for a time shifted towards more gene-based approaches. In these so called reverse genetic analyses, interesting genes are identified thanks to various methods such as evolutionary conservation or expression in response to stimuli. The function(s) is(are) then determined owing to gene inactivation, as depicted in the chapter "Methods for macromolecules extraction and genetic transformation". However, with the development of the next generation sequencing (NGS) technologies, gene and mutation identification through complete sequencing of mutant genome have recently been back to the fore, reinvigorating the classic genetic analyses, i.e., the search for mutants affected in

<figure><img src="image_67.png" /><figcaption>**Figure 61** PM154: structure of the mating type locus and phenotype.</figcaption></figure>
<figure><img src="image_68.png" /><figcaption>**Figure 62** Schematic representation of the *P. anserina* circular mitochondrial genome as defined in 1990. This genome is a composite of the genome of strain s (small s) with three additional sequences (optional introns) from strain A.</figcaption></figure>

defined phenomena and the subsequent identification of the involved genes. Nowadays, genetic analyses rely on both methods, depending on the needs. Importantly, all these genetic methods require the availability of a high-quality sequence of the *P. anserina* genome and more recently on microarray data or
RNAseq transcriptomes for expression-based selection of candidate genes.

In fact, *P. anserina* was one of the first organisms to enter the genomic era, even before it was fashionable. Indeed, as early as 1990, the complete sequence of its circular mitochondrial genome was available. At that time it was the longest contig present in the GeneBank/EMBL/DDBJ databases! The sequence has been determined by the Maxam and Gilbert method for the complete 91 192-bp genome of strain S (small s), as well as for three additional sequences present in strain A with a genome of 100 314 bp (figure 62). Those who do/did sequencing with the Maxam and Gilbert method can truly appreciate the efforts engaged in determining this sequence! The more recent releases of the genome sequence of strain S (BIG S) and s (small s) established with NGS data largely confirmed the sequence obtained then. However, the new sequences contain about 70 differences with the old one. Note that the new mitochondrial sequence of strain S (BIG S) has only one polymorphism with that of strain s (small s), indicating that the differences with the 1990 version are not due to *de novo* mutations during domestication but rather are errors in the old sequence. Presently, annotation of the genomes of both strains (Table 6) defines the following genes: the large (*rnl*) and small (*rns*) precursor of the ribosomal RNA (rRNA), 27 functional and one non-functional tRNA enabling to decode mRNAs using a non-standard genetic code in which UGA is not a stop codon but a tryptophane (W) sense codon, and 53 protein coding genes. In addition, there is a remnant in the genome of an integrated copy of the pAL2-1 plasmid. The free linear pAL2-1 plasmid is present in some strains of *P. anserina*, but not in S and s. Among the 53 protein coding genes some may not be actual genes and most proteins are intron-encoded proteins involved in intron splicing, as is customary in many fungal mitochondrial genomes. The proteins involved in the respiratory pathway are cox1, cox2, cox3, cob, NAD1, NAD2, NAD3, NAD4, NAD4L, NAD5, NAD6, ATP6 and ATP8. This set is typical to what is found in related species. There is also a ribosomal protein (rps3) encoded within an intron of *rnl*.

Additionally, the nuclear genome sequence of *P. anserina* was one of the first available. Decision to establish the sequence was made in 1999 and the first proposal deposited as an answer to a call by Genoscope, at that time the main agency for genomic analysis in France, at the beginning of the year 2000. Because establishing the complete genome sequence appeared an unsurmountable task for the small community working with the friendly mold without additional information, the proposal aimed first at establishing the genome sequence around the centromere of chromosome 5 of strain S. The project was accepted, provided that the DNA came from bacterial artificial chromosomes (BAC) and not cosmids
<table><caption>Table 6: main features of strain S genome</caption><thead><tr><th>number of chromosomes</th><th>7 nuclear + 1 mitochondrial</th><th>the number of chromosome is as expected similar to the one determined by classical genetics (see map figure 54)</th></tr></thead><tbody><tr><td colspan="3">genome size (nuclear + mitochondrial)= 36.06 Mb + 94.2 Kb</td></tr><tr><td>chromosome 1</td><td>~8 838 400</td><td>7 gaps remain on this chromosome; the left telomere sequence is not available</td></tr><tr><td>chromosome 2</td><td>~5 260 700</td><td>4 gaps remain on this chromosome; the right telomere sequence is not available</td></tr><tr><td>chromosome 3</td><td>~4 736 000</td><td>the rDNA locus is present in a sub-telomeric position of the left arm; estimation of the number of rDNA repeats is 70; 3 gaps remain on this chromosome</td></tr><tr><td>chromosome 4</td><td>~3 887 200</td><td>7 gaps remain on this chromosome</td></tr><tr><td>chromosome 5</td><td>~4 864 800</td><td>2 gaps remain on this chromosome; the left and right telomere sequences are not available</td></tr><tr><td>chromosome 6</td><td>~4 300 800</td><td>no internal gap remain on this chromosome, but the right telomere sequence is not available</td></tr><tr><td>chromosome 7</td><td>~4 174 400</td><td>11 gaps remain on this chromosome; the right telomere sequence is not available</td></tr><tr><td>mitochondrial chromosome</td><td>94 198</td><td>the genome is circular</td></tr><tr><td colspan="3">nuclear gene number: protein coding + tRNA + 5S rRNA (pseudogenes)= 10757 (434) + 361 (5) + 87 (8)</td></tr><tr><td>chromosome 1</td><td>2711 (84) + 66 (1) + 23 (2)</td><td></td></tr><tr><td>chromosome 2</td><td>1618 (53) + 42 (2) + 17 (2)</td><td></td></tr><tr><td>chromosome 3</td><td>1232 (52) + 49 (1) + 13 (2)</td><td></td></tr><tr><td>chromosome 4</td><td>1158 (45) + 50 (0) + 9 (1)</td><td></td></tr><tr><td>chromosome 5</td><td>1470 (116) + 63 (0) + 8 (0)</td><td></td></tr><tr><td>chromosome 6</td><td>1321 (52) + 44 (0) + 12 (1)</td><td></td></tr><tr><td>chromosome 7</td><td>1247 (32) + 47 (1) + 12 (1)</td><td></td></tr><tr><td colspan="3">mitochondrial genes: 53 protein coding genes + 27 tRNA + 1 tRNA pseudogene + small rRNA + large rRNA</td></tr></tbody></table>

(which required the construction of the BAC library and its screening for centromere of chromosome 5 DNA). About 480 kb of sequence was established and annotated. The paper reporting the data was published in 2003. This success, as well as decreases in sequencing costs and introduction of better informatics programs for genome sequence assembly, led to the proposal at Genoscope to establish the complete sequence of strain S *mat+* by sequencing the extremities of plasmids having 3 kb or 12 kb inserts, as well as those of BACs and cosmids, with the Sanger methods using fluorescent dyes. To facilitate annotation, the sequences of about 50 000 cDNAs was also established. All the data were made available through internet as early as the end of 2006 and the official paper was published in 2008. Since then, the genome assembly and accuracy of strain S *mat+* has been refined by 454 and Illumina NGS data. The annotation has also been improved thanks to RNAseq information from many developmental stages of the fungus (one-day-old and four-day-old mycelia, two-day-old and four-day-old perithecia, non-germinated ascospores and eight-hour-old germinated ascospores, see "Methods for macromolecules
composed of AT-rich sequences originating from transposons, likely through the accumulation of RIP

<figure><img src="image_69.png" /><figcaption>**Figure 63** Schematic representation of the *P. anserina* genome as defined by complete sequencing. The chromosome are to scale (note the very small size of the mitochondrial chromosome). The blue dots are the centromeres. Main genetic markers are indicated on the left side, including classical genetic markers (i.e., mostly the mutations generated in strains S or s) and newly-constructed gene deletions. On the right side are the polymorphisms with strain T used to help with genome assembly. The rDNA is on the left arm</figcaption></figure>
of chromosome 3 in a sub-telomeric position. The purple line identifies the region devoid of recombination around the mating type.
mutations. RIP is a phenomenon first described in *N. crassa* and later on in *P. anserina*. This phenomenon results in the accumulation of C to T mutations in duplicated sequences. Only sequences duplicated in the same nucleus are inactivated, while those present in single copy in the other nucleus are not affected.
The mutations may thus be prompted during the dikaryotic stage of the sexual tissues. While very efficient in *N. crassa*, the phenomenon is less pronounced in *P. anserina*, since only ascospores produced late harbor mutations and the mutation frequency is low (0.3 -with unlinked repeats- to 15 –with tandem repeats- mutations per kb were detected by sequencing). Nevertheless, duplicated sequences such as transposons accumulate C to T mutations through multiple rounds of sexual reproduction, resulting in high AT content. In the centromere, this percentage is about 75%. Centromeres measure from 56.5 kb for chromosome 3 to 87.5 kb for chromosome 2. Chromosome 1 centromere may be bipartite. This chromosome is twice as big as the others (Table 6, figure 63) and may have resulted from the fusion of two acrocentric chromosomes by their centromere-carrying ends. Indeed, there are two large AT-rich regions of 81 kb and 38 kb separated by a 500-kb region devoid of recombination. The 81-kb region has been assigned the role of centromere, but lack of recombination between the two regions may be indicative that the 38-kb one may also play a role as centromere.

Telomeres are also typical for eukaryotes. They end by GGGTTA repeats. Although not all ends are properly assembled, there are sequence reads containing these repeats necessary for all chromosome ends being capped this way. In sub-telomeric regions, there are also transposons inactivated by RIP. These regions have variable sizes, ranging from 2-3 kb to several dozens of kb. They often contain a degenerate helitron, as described in other fungi. Transposons, often associated with small segmental duplications, are present within the chromosome arms. They are mostly nested together in regions that may reach sizes up to 50 kb. RIP has also acted on these regions, which are thus rich in AT. There are at least 51 different transposons (Table 7), few being functional due to inactivation by RIP. The largest is the retrotransposon "Grenouille", measuring nearly 12 kb, and the smallest is the retrotransposon "Eleutherodactylus", which is only present as LTR (Long Terminal Repeat) of 58 bp. Note that many repeated sequences are of unknown affinity (Table 7). Annotation of these sequences is ongoing, since assembly of the repeated regions still contains gaps. Overall, these repeated regions cover about 5-10% of the genome.

Other repeated regions are the ones coding for the rRNA. There are 87 copies of the 5S rDNA
gene scattered all over the genome (Table 6) and the rDNA locus, which is in a sub-telomeric position on

<table><caption>Table 7: *P. anserina* strain S transposons and repeated sequences of unknown affinity</caption><thead><tr><th>Family</th><th>Name</th><th>Element size (bp)</th><th colspan="3">Estimated copy number</th></tr><tr><th>DNA transposons</th><th></th><th></th><th colspan="3">LTR size</th></tr></thead><tbody><tr><td rowspan="5">mutator-like</td><td>dendrobates</td><td>3 449</td><td>65</td><td colspan="2">3</td></tr><tr><td>discoglosse</td><td>1 867</td><td>49</td><td colspan="2">153</td></tr><tr><td>pelobate</td><td>1 948</td><td>45</td><td colspan="2">20</td></tr><tr><td>rainette</td><td>1 298</td><td>41</td><td colspan="2">9</td></tr><tr><td>scaphiopus</td><td>1 428</td><td>23</td><td colspan="2">1</td></tr><tr><td rowspan="3">Tc1/mariner-like</td><td>altiphrynoides</td><td>403</td><td>50</td><td colspan="2">5</td></tr><tr><td>discodeles</td><td>257</td><td>40</td><td colspan="2">49</td></tr><tr><td>mantella</td><td>383</td><td>46</td><td colspan="2">18</td></tr><tr><th>LTR retrotransposons</th><th></th><th>full element</th><th>LTR</th><th>full element</th><th>solo LTR</th></tr><tr><td rowspan="3">copia/Ty1</td><td>centrolene</td><td>5 918</td><td>182</td><td>18</td><td>14</td></tr><tr><td>hyla</td><td>6 466</td><td>630</td><td>10</td><td>63</td></tr><tr><td>nephelobates</td><td>5 594</td><td>445</td><td>7</td><td>0</td></tr><tr><td rowspan="5">gypsy/Ty3</td><td>crapaud</td><td>7 372</td><td>400</td><td>80</td><td>&gt;300</td></tr><tr><td>grenouille</td><td>11 866</td><td>360</td><td>20</td><td>53</td></tr><tr><td>pipa</td><td>&gt;4 242</td><td>?</td><td>1</td><td>0</td></tr><tr><td>yeti</td><td>6 943</td><td>355</td><td>2</td><td>37</td></tr><tr><td>boophis</td><td>?</td><td>367</td><td>0</td><td>3</td></tr><tr><td rowspan="11">solo LTR</td><td>alytes</td><td>?</td><td>230</td><td>0</td><td>3</td></tr><tr><td>arthroleptis</td><td>?</td><td>238</td><td>0</td><td>10</td></tr><tr><td>ascaphus</td><td>?</td><td>170</td><td>0</td><td>6</td></tr><tr><td>astylosternus</td><td>?</td><td>172</td><td>0</td><td>12</td></tr><tr><td>atelopus</td><td>?</td><td>601</td><td>0</td><td>9</td></tr><tr><td>bufoides</td><td>?</td><td>153</td><td>0</td><td>6</td></tr><tr><td>capensibufo</td><td>?</td><td>256</td><td>0</td><td>11</td></tr><tr><td>cochranella</td><td>?</td><td>155</td><td>0</td><td>18</td></tr><tr><td>crepidophryne</td><td>?</td><td>137</td><td>0</td><td>6</td></tr><tr><td>cyclorana</td><td>?</td><td>194</td><td>0</td><td>8</td></tr><tr><td>dendrophryniscus</td><td>?</td><td>73</td><td>0</td><td>5</td></tr><tr><td>eleutherodactylus</td><td>?</td><td>58</td><td>0</td><td>14</td></tr><tr><td>nyctibates</td><td>?</td><td>169</td><td>0</td><td>5</td></tr><tr><td>pelodytes</td><td>?</td><td>128</td><td>0</td><td>8</td></tr><tr><td>rana</td><td>?</td><td>508</td><td>0</td><td>17</td></tr><tr><td>xenopus</td><td>?</td><td>235</td><td>0</td><td>16</td></tr><tr><th>helitron</th><td>epipadobates</td><td>8388</td><td colspan="3">22</td></tr><tr><th>unknown affinity</th><td></td><td></td><td colspan="3"></td></tr><tr><td></td><td>adenomus</td><td>382</td><td colspan="3">14</td></tr><tr><td></td><td>alloporyne</td><td>268</td><td colspan="3">9</td></tr><tr><td></td><td>amolops</td><td>585</td><td colspan="3">31</td></tr><tr><td></td><td>andinoporyne</td><td>122</td><td colspan="3">12</td></tr><tr><td></td><td>ansonia</td><td>90</td><td colspan="3">5</td></tr><tr><td></td><td>batrachyla</td><td>192</td><td colspan="3">20</td></tr><tr><td></td><td>bombina</td><td>680</td><td colspan="3">10</td></tr><tr><td></td><td>bombinator</td><td>472</td><td colspan="3">5</td></tr><tr><td></td><td>brachycephalus</td><td>259</td><td colspan="3">5</td></tr><tr><td></td><td>bufo</td><td>699</td><td colspan="3">9</td></tr><tr><td></td><td>callulops</td><td>302</td><td colspan="3">5</td></tr><tr><td></td><td>cardioglossa</td><td>182</td><td colspan="3">5</td></tr><tr><td></td><td>churamiti</td><td>143</td><td colspan="3">6</td></tr><tr><td></td><td>lechriodus</td><td>246</td><td colspan="3">13</td></tr><tr><td></td><td>leptodactylodon</td><td>570</td><td colspan="3">22</td></tr><tr><td></td><td>leptopelis</td><td>112</td><td colspan="3">6</td></tr><tr><td></td><td>microhyla</td><td>1179</td><td colspan="3">8</td></tr><tr><td></td><td>platymantis</td><td>593</td><td colspan="3">8</td></tr><tr><td></td><td>schoutedenella</td><td>241</td><td colspan="3">7</td></tr></tbody></table>
the left arm of chromosome 3. Based on the frequency of sequence reads for the rDNA versus that of single copy regions, the number of copy of the rDNA has been estimated to 70, which is in line with the 87 copies for the 5S genes. The rDNA unit measures 8 083 pb and is organized classically: 18S precursor – ITS1 – 5.8S precursor- ITS2 – 28S precursor- large intergenic region. Note that the large intergenic region contains 3 different small repeated sequences of about 40, 100 and 150 bp.

On figure 63, in addition to the polymorphic markers used to construct the linkage map used for assembly are indicated some of the available genetic markers that can be used for genetic mapping. These results from both forward genetic mutants for which the affected gene has been identified and reverse genetic mutants. As seen, the genome, including the mitochondrial chromosome, is well covered. Important markers are firstly the mating type, with its two idiomorphs *mat+* and *mat-*, located on chromosome 1. As exposed above, this locus has been deleted and the resulting strain is used in developmental genetic analyses (see "Developmental genetics: grafting and genetic mosaics"). A chimaeric *mat+ mat-* locus is also available in strain PM154, a strain also used to study fruiting body development (see also "Developmental genetics: grafting and genetic mosaics"). The second important marker is the mid26 one. It is located in the mitochondria and is the clean elimination of the first intron of the *cox1* gene. This intron is involved in the shortening of the life span of the fungus. Without the intron, strains live three times longer! Introducing this marker into short-live strains make them more amenable to analysis. First obtained in the s (small s) strain, this marker has since been introgressed into the S (BIG S) genetic background by 10 backcrossed with the S strain. Introduction is simply done by crossing the desired strain as male to the mid26 strain as female. All progeny will carry the mid26 marker.
# Part 2: *Podospora anserina* as a model for physiological and molecular analysis
In this second part of the book, several features of *P. anserina* lifecycle and lifestyle will be introduced, especially those that involves analysis of molecular pathways. Note that studies of these have often led to a better understanding of biology. Indeed, *P. anserina*, owing to its ease of use and speed of the sexual cycle, is an excellent model to tackle problems that would be hard to study with other organisms such as ageing or recognition of self-versus-non-self. However, the smallness of the community working with this fungus makes it such that our knowledge on many subjects is often patchy. We shall see in turn all aspects of the *P. anserina* stages: spore, vegetative thallus, namely the mycelium and finally the reproductive phase.
Ascospore ejection and germination

As the first stage of the lifecycle of the fungus, ascospores have two roles: dispersal and persistence. Surprisingly, modalities of both phenomena are poorly known for *P. anserina*. As previously stated, ascospore are forcibly discharged out of the fruiting body thanks to a turgor pressure-operated mechanisms. The deletion of the *Trk-1* gene involved in potassium import inside the cell impairs this process, while deletion of the genes involved in export results also in lack of spore ejection on medium with high potassium concentration. This suggests that proper turgor driven by intracellular potassium concentration is essential for correct ascospore ejection. Asci are ejected one at the time out of the fruiting body and the four (or five) ascospores often stick together thanks to the actin rope that links spores. Ejection distance is a few dozen centimeters and it is probable that wind also participate in later stages of dispersal before asci land on neighboring plants. How long in the wild ascospores may persist is not known, but it is likely that their lifespan outside the fruiting bodies is short. Indeed, in the lab we know that ascospores kept a few weeks at 4°C on agar covers fail to germinate and that their drying results in complete failure of further germination. However, we also know that *P. anserina* is able to resist in the wild on dried dung for a long time (several weeks) and renew rapidly ejection of viable ascospore when rehydrated. Yet, on Petri plates, ascospore production and ejection is irreversibly stopped if the conditions are too dry or two cold. Finally, we know that drying fertile mycelium with fruiting body-carrying ascospores overnight at 65°C does not result in death of the fungus. From all of these scattered data, the best guess is that ascospore ejection is sensitive to drying but that somehow natural drying; which may be slower that what is done in the lab, may result in preservation of the ejection function. Also, ascospores drying inside the fruiting body may survive, while they are much more fragile after ejection.

In the wild, the next step for ascospores is to be ingested by an herbivore to trigger germination. This stimulus is complex as it involves heat shock (37°C is a lethal temperature for the fungus, while 36°C is not...), drastic pH change that may differ from one type of animal to the other, attack by proteases, carbohydrate hydrolases and biliary salts... While some fungi require an exact combination of these for
germination, *P. anserina* is not demanding in its germination conditions, accounting for its presence not only in dung but in other biotopes. Indeed, while the G medium is routinely used to obtain 100% germination, high percentage of germination (often >80%) can be obtained by treating ascospores with heat shock (55-65°C for ~30 minutes), H₂O₂ (3% for 20 minutes), phenolic compounds (10⁻³-10⁻⁴ M) and action of pancreatin (1-2 h at 37°C). It is likely that many other as yet untested stresses can trigger germination of *P. anserina* ascospores.

The molecular pathway involved in the triggering of germination has been addressed by

<figure><img src="image_70.png" /></figure>

**Figure 64** Schematic representation of the *P. anserina* ascospore germination pathway. Germination is triggered by several stresses that may act through different and as-yet unknown receptors. These activate through unknown mechanisms the PaMpk2 MAPK kinase pathway, which in turn activates the PaNox2/PaPls1 NADPH oxidase. Activation of this complex results in reactive oxygen species (ROS) production, likely superoxide anions. This promotes the breaching of the germination pore by the germination hyphae and germination of the ascospore.
identifying mutants impaired in germination. These are affected in the PaNox2 NADPH oxidases, the PaPls1 tetraspanin and the PaMpk2/PaMKK2/PaTLK2 MAP kinase module. In these mutants, ascospore germination is completely abolished. Conversely, a mutant having a constitutively active PaMpk2 MAP kinase germinates spontaneously at high frequency. Introducing the constitutive mutation into the PaNox2 and PaPls1 mutant background fails to restore germination suggesting that the MAPK module acts upstream of PaPls1 and PaNox2. A model presenting the germination pathway in *P. anserina* is proposed in figure 64. Pigmentation is also an important factor in the process since lack of pigment, effected either by genetic or chemical inactivation of the melanin pathway (see next chapter) results in spontaneous germination, even in the presence of the PaNox2, PaPls1 and PaMpk2 mutations. Other mutants presenting ascospore germination defects are known, especially those affected in peroxisomal proteins. However, peroxisomal mutants also present ascospore pigmentation defects, making it difficult to know whether germination impairment is a direct consequence of the mutation or an indirect one linked to pigmentation problems. Interestingly, a connection between ascospore germination and development of appressorium-like structures involved in biomass penetration has been found. Indeed both processes are under the same genetic control (see next chapter).

As explained in the chapter on morphology, *P. anserina* is able to produce sometime in very large amount spermatia, questioning their possible role as dispersal units. Cytological analysis revealed that spermatia contain one nucleus and a mitochondrion embedded inside a fair volume of cytoplasm. They carry thus everything needed to germinate. Yet all attempts, but one that could not be reproduced recently, made to trigger germination failed. Even in the successful case, only 1% germination was achieved. Pending the discovery of efficient conditions for germination, spermatia are thus best considered as serving only as male gametes.
Mycelium growth and development

Owing to the small sizes of the hyphae, *P. anserina* is not routinely used to study the basic
mechanisms involved in mycelium growth, unlike *N. crassa* in which many studies are made. I will thus
rather emphasize phenomena occurring during the vegetative phase, namely, Senescence, Crippled
Growth and other phenotypic instabilities, vegetative/heterokaryon incompatibility including its control
by the Het-s prion, hyphal interference and differentiation of appressorium-like structures. I will start
with some basic description of the physiology and metabolism of the mycelium.

Growth of the fungus is best achieved on medium with a pH=7.2-7.6, which is the pH value of the M2 medium. After growth the pH is still equal to about 7, indicating that *P. anserina* does not drastically alter the pH of its growth medium. The usually-used temperature for cultivation is 27°C. Lethal temperatures are over 36°C. In many strains, as previously stated in Part 1, there is a polymorphic gene closely linked to the mating type that influences growth at high temperature. In many strains, including strain S, the *mat*- isolates grow better than *mat+* ones at high temperature. For example at 36-37°C, the S *mat*- isolates differentiate aerial hyphae, while the S *mat+* ones do not. This gene also controls longevity (see below). As expected, the fungus grows slowly at low temperatures and it still grows at 10°C. Light is not necessary for growth but is required for proper pigmentation. Major pigments are melanin synthesized by the dihydroxynaphtalene (DHN) pathway. The PKS acting at the first stage of the pathway has been identified and is required for pigmentation at all stages of the lifecycle (mycelium, fruiting bodies and ascospores). On the contrary, only the laccases acting in the ascospores have been identified. These enzymes are involved at the last stage of pigment production, i.e., during the polymerization of the 1-8-dihydroxynaphtalene precursors into melanin inside the cell wall. Hyphae need to be about two-day-old to start to accumulate pigment. In the absence of melanin, the mycelium often accumulates after a few days a pinkish color, likely resulting from the oxidation of cytochromes.

The fungus is able to consume various carbon sources to promote mycelium growth, but not all of them may sustain further sexual development. Indeed, glucose is efficiently utilized for vegetative
growth but results in severely diminished fertility, when compared to dextrin, starch or amorphous
cellulose (paper). Hence, the fungus is almost sterile on potato dextrose agar (PDA). Noteworthy, P.
*anserina* lacks invertase and is thus unable to consume sucrose. Intriguingly, glycerol is also not
metabolized well, although *P. anserina* possesses the transporters and enzymatic pathway to metabolize
this compound. The fungus thrives on complex biomass, including dried hay, straw, miscanthus, oat meal,
cotton seed hulls, soybean hulls and many woody substrates. Dung is of course well used; however, we
see some fluctuation in the ability of the fungus to produce mycelium and fruiting bodies on different
dung pellets, even from originating from the same animals. Growth requires oxygen since the fungus
does not propagate, but does not die, in anaerobiosis. Nitrogen also has a large impact on growth and
fertility. The fungus is for example sterile on media containing more than 1 g/L of yeast extract. It is also
sterile in the presence of too much ammonium ions. The fungus is thus routinely grown on autoclaved
urea, but it can also use nitrate and various amino acids. It requires thiamin for growth and both thiamin
and biotin for fertility (see below the section on sexual reproduction). In summary, *P. anserina* can grow
on many media, but is rather fussy when it comes to sexual reproduction, then the growth medium must
not be too rich in both carbon and nitrogen sources, and must contain biotin. Please, see below the
section on "Fruiting body production and repartition" in the chapter on "Sexual Reproduction" for
detailed analysis of the parameters enabling sexual reproduction.

All the above parameters influence the morphology of the mycelium that may or may not follow
rhythms. Rhythmic growth is also under a genetic control since many mutants having such pattern of
growth on all media have been described.

Senescence and the Premature Death Syndrome

Unlike what is observed in many fungi, the vegetative growth of *P. anserina* is not unlimited. It stops definitively at one point and is accompanied by the death of the apical cells (figure 65). This phenomenon is called Senescence and is used as a simple model to study ageing. In fact, despite the claim by many that senescence was first observed on human fibroblasts, limited cell division was first described in *P. anserina* as the publication dates of the papers attest (1953 for *P. anserina* and 1961 for fibroblasts). To date, all strains taken from nature that belong to one of the seven species of the *P*.
<figure><img src="image_71.png" /><figcaption>**Figure 65** Senescence in *P. anserina*. On this culture grown in race tubes, ascospore germination occurred on the left. The mycelium then developed toward the right until growth ceased. Intense pigmentation or "Barre de Sénescence" accompanies the growth stoppage. In Barre de Sénescence, cells are dead. Cells located closer from the germination point are still alive. This culture grew for about three weeks, i.e., for 10 cm, before dying.</figcaption></figure>

*anserina* species complex have been observed to undergo Senescence, albeit with different life span that may range from 2-3 centimeters to more than 20 cm on M2 medium, indicating that Senescence is a constant features of these fungi.

At the cellular level, Senescence is accompanied by a drastic reorganization of the internal
membrane networks (mitochondria and the nuclear membrane/endoplasmic reticulum network). Often
the fungus is female-sterile close to the "Barre de Senescence", although few and abnormally-maturing
perithecia may sometime be differentiated. Male fertility appears not drastically affected. At the
molecular level, Senescence is correlated with mitochondrial DNA modifications (figure 66). The one that
is the most often observed is the accumulation as a circular plasmid deriving from the first intron of the
*COX1* genes. This DNA molecule is called senDNAα or pIDNA. Nevertheless, other parts of the
mitochondrial DNA may amplify as circular molecules during Senescence, the senDNAβ and senDNAγ.
Intriguingly, the PAL2-1 linear plasmid hosted by the mitochondria of some strains promotes long life
(similar plasmids are known to trigger senescence-like processes in *Neurospora spp*.). Early formal
analyses have shown that Senescence is caused by the accumulation of a cytoplasmic and infectious
element, called the "determinant of Senescence" and whose molecular nature is still unknown. This was
demonstrated by several experiments, including formal analysis of the longevity of subcultures (figure
67), analysis of the segregation patterns of senescence (figure 68) and micromanipulation of hyphae
under the microscope (figure 69). The relationship between the mitochondrial DNA modifications and the
cytoplasmic and infectious determinant is not clear, since they were shown not to co-segregate both
<figure><img src="image_72.png" /><figcaption>**Figure 66** Mitochondrial DNA modifications observed during Senescence in *P. anserina*. Left, scheme of the mitochondrial DNA in young cultures and origins of the senDNA molecules amplified during Senescence. Right, agarose gel stained with ethidium bromide showing the structure of the mitochondrial DNA molecules in young (Y) cultures and in senescent ones (S1 & S2). In S1, the amplified molecule is senDNAα while senDNAα is not amplified in S2. MW: molecular weight.</figcaption></figure>

during contamination experiments and meiosis. Especially, senDNAα, which was thought at one point to be the determinant of Senescence, does not accumulate in some senescing strains. Moreover, a strain deleted for the first intron of COX1, and thus unable to amplify senDNAα, was found to undergo Senescence, albeit with a longer lifespan.

Longevity can be easily monitored with *P. anserina* since it can be measured in centimeters of growth! There is thus no need for constant surveillance of the culture to accurately measure longevity, a feature that is not possible in many models, especially in animals that need constant surveillance to determine the actual time of death. Longevity is under the control of numerous nuclear genes,
<figure><img src="image_73.png" /><figcaption>**Figure 67** Lifespan analysis in *P. anserina*. Top, lifespan curves of two strains, one short-lived and the other long-lived. The logarithm of the percentage of surviving cultures is given as function of the distance between the inoculation point and the Barre de Senescence. These curves have two different parts. The first one in which the percentage is equal to 100 % shows that all cultures are able to grow for certain duration, called the incubation period. The second one is typical of the occurrence of a random event with a constant probability (given by the slope of the line). Interpretation of these curves is given below. In cultures, a random event occurs, the commitment to Senescence (i.e., the appearance of the Determinant of Senescence; red S). Once this determinant has appeared, it amplifies exponentially following a constant kinetics before it kills the cells. This model can be verified by re-inoculating explant to new medium: those taken after commitment yields short-lived cultures, while those taken before commitment will generate various lifespans following the same curve as the first cultures.</figcaption></figure>
<figure><img src="image_74.png" /><figcaption>**Figure 68** Segregation analysis of Senescence in *P. anserina*. Senescence is transmitted to the progeny only when the maternal parent is senescent showing that the Determinant of Senescence is located within the cytoplasm. When transmitted during ascospore formation, senescent thalli resulting from the germination of ascospore carrying the determinant are very short-live (< 1-2 cm). Their percentage increases exponentially as the maternal parent is nearer to the Barre de Senescence, confirming that Senescence results from the exponential accumulation of a cytoplasmic factor.</figcaption></figure>

mitochondrial ones and some less defined cytoplasmic genetic elements. It is also highly sensitive to environmental conditions and various chemical compounds. We shall now see in turn each of these parameters, starting with the role of nutrients and chemicals.

Composition of the growth medium greatly affects longevity. For example, longevity is three to four times shorter on M2 than on Corn Meal (MR) medium. Paramount is the quantity of carbon source since caloric restriction delays all symptoms of Senescence, but the nitrogen source is also important (for example presence of yeast extract alters the life span, but in a complex manner depending upon the concentration). Longevity is also dependent upon temperature with the hotter the incubation, the shorter the lifespan. Low temperature can even trigger rejuvenation, as do surprisingly prolonged incubation at high temperature and many treatments that inhibit growth such as incubation in distilled water, storage under oil or incubation under nitrogen air. Unexpectedly, light shortens lifespan by an unknown mechanism.

Presence of many chemical compounds in the growth medium also affects the life span. I shall
<figure><img src="image_75.png" /><figcaption>**Figure 69** Micromanipulation analysis of Senescence in *P. anserina*. Young and senescent hyphae are brought together under a microscope as to observe anastomosis formation, which proceeds after a few hours. Once anastomoses are formed, the hyphae are incubated for 10 hours. Hyphae continue their growth, especially the young ones. At the end of the incubation, the formerly young hyphae are fragmented with a micromanipulator and new cultures are generated from the isolated fragments. Longevity analysis shows that fragments taken close to the anastomosis are invariably senescent, while those farther away are more or less senescent or even still young, demonstrating the infectious nature of the Determinant of Senescence.</figcaption></figure>

only emphasize those that inhibit the respiratory chain that increase life span up to near-immortalization especially chloramphenicol and ethidium bromide that can cure senescence and rejuvenate the culture. Ethidium bromide was the most studied compound and it was shown that its action result in the elimination of the Determinant of Senescence as well as the restoration of an intact mitochondrial DNA. Contrary to inhibitors of mitochondrial functions, inhibitors of cytosolic translation, especially cycloheximide, shorten lifespan.

Genetic determinants of lifespan have also been thoroughly investigated. The first genetic determinant discovered by G. Rizet was a nuclear one called *incolore* (also known as *incoloris*). *Incolore* mutants have pleiotropic defects, including low pigmentation, profuse aerial hyphae and spermatia, female sterility and longer lifespan. To date, the affected gene in the first identified *Incolore* mutant, *IncA*, has not been identified. I will further discuss the incolore mutants later on this chapter. *Incolore* is only one of the many mutants modifying lifespan. Indeed, a survey of known mutants and of strains generated by insertional mutagenesis, the first one to be performed on any organism, indicated that between 10 and 50% of the genes likely control lifespan, *i.e.*, between 1000 and 5000 genes. Among those, genes
involved in mitochondrial biology and in cytosolic translation appear the most important.

Firstly, mutants affected in the respiratory chain have increased lifespan. Overexpression of the alternative oxidase, an enzyme able to replace the cytochrome C oxidase, in these mutants restore a normal lifespan along with a more efficient respiration and lower reactive oxygen species production. When the mutations are present in mitochondrial genes, effect on longevity can be tremendous and many mitochondrial mutants affected in genes encoding subunits of the respiratory chain are immortal. At the present time, it is not clear how the activity of the respiratory complexes controls Senescence, through metabolism, through reactive oxygen species or other processes. Indeed, mutants often have pleiotropic defects, including retrograde effects on nuclear gene expression, making it difficult to disentangle their actions on lifespan. Mitochondrial fusion and fission, mitophagy and other autophagy pathways, mitochondrial import, protein quality control and many other processes related to mitochondrial physiology (e.g., reactive oxygen species production and removal, programmed cell death...) have been shown to control lifespan. It is thus likely that each mutation involved in mitochondrial biology alters several lifespan-prolonging and lifespan-reducing pathways. It is beyond the scope of this book to present in detail all the researches carried out on the mitochondria roles in Senescence and longevity control in P. anserina, which is still a very active field of research. There are several reviews on the subject referenced in the appropriate section of this book.

Secondly, cytosolic translation, the main energy consumer of the cell, is also a major actor of
lifespan control, linking again energy metabolism with ageing as in other organisms (e.g., *Caenorhabditis elegans*). Some translation mutants may even be immortal without any obvious additional physiological
defect. Indeed, mitochondria-related mutants, especially the immortal ones affected in mitochondrial
genes, are slow and spindly growing, lack pigmentation and aerial hyphae, and are female sterile. On the
contrary, I have obtained an immortal strain having no obvious defect during growth of the mycelium
(same speed, color morphology, etc.) and sexual reproduction (fruiting bodies are differentiated in similar
amount and with the same kinetics as in the wild type). This strain carries two different alleles of the
genes encoding the cytosolic translation elongation factor eEF1A. Once again, how cytosolic translation
affects Senescence and lifespan is not understood and its action may be through modification of several
lifespan-prolonging and lifespan-reducing pathways. An additional very interesting feature of the
translation mutants, especially those modifying its accuracy, is that they change the spectrum of the
mitochondrial DNA modification observed in dying hyphae. Indeed, some have impaired amplification of
<figure><img src="image_76.png" /><figcaption>**Figure 70** Genesis of the deletion amplified during the Premature Death Syndrome in *P. anserina*. Following transposition of the class II intron α (the first intron of the *COX1* gene), two copies are present in the same orientation in the mitochondrial genome. Such molecule can be detected by PCR amplification in all tested strains. A recombination between the two copies yields two molecules. Only the large one amplifies during the Premature Death Syndrome.</figcaption></figure>

the senDNAα and thus carry exclusively other amplified molecules belonging to the senDNAβ and senDNAγ families (see S2 in figure 66). The most interesting one is the *AS1-4* mutation affecting the *AS1* gene encoding the S12 protein of the small ribosomal subunit. Indeed, this mutation triggers a near immortality in *S mat+* cultures and a very short lifespan in *S mat-* ones (1-2 cm). This latter short lifespan has been coined as “Premature Death” owing to the fact that the DNA modification observed in dying hyphae is unique and not seen during normal Senescence. It consists in the deletion of one third of the mitochondrial DNA (figure 70). Interestingly, one of the borders of the deletion is the intron α amplified
as senDNAα during senescence. It has been shown that this intron is also a transposon that integrates
near the tRNA inserting serine during mitochondrial translation. The deletion amplified during Premature
Death is the result of an intramolecular recombination between the two copies of the intron/transposon
(figure 70). Why only one of the two molecules generated by the recombination (figure 70) is amplified is
unknown. Interestingly, senDNAα is likely generated in a similar manner. Transposition of intron α just
upstream itself could yield a tandem duplication of the intron. A recombination between the two copies
can then generate the first extrachromosomal circular copy of senDNAα, which can subsequently be
amplified either by the same mechanism or by replicating. Finally, the genetic determinant linked to the
mating type that modulates the lifespan difference between the *mat-* and *mat+* isolate has been
identified. It is the *rmp1* gene that contains a polymorphism between the *mat+* and *mat-* in some, but not
all, natural isolates. This polymorphism also controls thermosensitivity (see above) and the long-known
difference in longevity between the *mat+* and *mat-* isolates in some strains.

Overall, a lot has been learned by studying Senescence in *P. anserina*, especially on the
physiology of respiration in connection with mitochondrial normal and abnormal functioning in an
obligate aerobic organism. Yet, owing to the complexity of the phenomenon, much remain to be
understood. With the advent of omics data, study of Senescence has now been upgraded to the study of
whole pathways instead of single genes, which should further deepen our understanding of ageing.

Crippled Growth and other phenotypic instabilities

Another cell degeneration process affecting the growth of *P. anserina* was discovered more
recently. Hyphae affected by this so-called “Crippled Growth” phenomenon grow more slowly, have
abnormal morphology and often accumulate high level of pigments. On the whole thallus, Crippled
Growth manifests as random-appearing sectors of extending altered growth with fewer and pigmented
hyphae (figure 71). In the sector, there is no aerial hyphae and the mycelium is female sterile. The
degeneration occurs only in *P. anserina* (but is present in all investigated strains) and not in its sibling
species, unlike Senescence. It also develops specifically on some media such as M2 supplemented with
yeast extract (figure 71), while wild-type strains of *P. anserina* never present Crippled Growth on the M2
medium. Like Senescence it is linked to the presence of a cytoplasmic and infectious factor (figure 72).
However, unlike Senescence this factor, called *C*, accumulate in stationary phase hyphae (figure 73).
<table><thead><tr><th>Growth regimen</th><th>M2</th><th>M2+YE</th><th>M2+ Ac. NH<sub>4</sub></th></tr></thead><tbody><tr><td>WT</td><td>NG</td><td>NG/CG</td><td>NG</td></tr><tr><td>IDC</td><td>NG</td><td>NG</td><td>NG</td></tr><tr><td>PDC</td><td>NG/CG</td><td>NG/CG</td><td>NG/CG++</td></tr></tbody></table><table><thead><tr><th>Presence of C in stat. ph.</th><th>M2</th><th>M2+YE</th><th>M2+ Ac. NH<sub>4</sub></th></tr></thead><tbody><tr><td>WT</td><td>+</td><td>+</td><td>+</td></tr><tr><td>« pink » IDC</td><td>-</td><td>-</td><td>-</td></tr></tbody></table>

**Figure 71** Crippled Growth in *P. anserina*. Left, a "Crippled Growth" sector develops on the uppermost thallus. Top right table, Growth regimen observed on different media for wild type (WT), mutants impaired for Crippled Growth (*IDC*) and those promoting Crippled Growth (*PDC*); NG: only Normal Growth occurs. NG/CG: Normal Growth with sectors of Crippled Growth. NG/CG++: sectors have a much altered growth as compared with M2+YE. Bottom right table, *C* is always present in wild-type (WT) mycelia during stationary phase and is absent from "pink" IDC mutants.

<figure><img src="image_77.png" /></figure>

**Figure 72** Contamination experiments showing that Crippled Growth is linked to the presence of a cytoplasmic and infectious factor. Left, a two-day-old Normal Growing thallus is inoculated with explants of Crippled Growing mycelium. After 5 days of incubation, Crippled Growing sectors develop downstream three out of the four tested CG explants. Marking the nuclei with the mating types indicates that CG sectors are of the recipient mating type, showing that Crippled Growth is due to the spreading of a cytoplasmic and infectious element.
<figure><img src="image_78.png" /><figcaption>**Figure 73** C is induced in stationary phase. Left, a 5-day-old Normal Growing wild-type culture on M2 is sliced as depicted by the two red rectangles. When inoculated on M2+YE, the slice gives a two-part cultures with on the right hyphae from the apical side of the slice regenerating Normal Growing hyphae and those on the left from the stationary phase side Crippled Growing ones. Note that on this plate the upper aerial part of the slice is on the top and the part that was at the bottom of the initial plate is at the bottom. The unequal shape of the Crippled Growth area of the new culture shows that C is induced earlier in the aerial part of cultures when compared with the bottom one. Contrary to what happens when the slice is inoculated on media permitting CG, inoculation on media not permitting CG, such as M2, yields only Normal Growing hyphae. Overall, this shows that C is induced in stationary phase hyphae even in conditions not permitting Crippled Growth, but will amplify in growing hyphae only in defined conditions, i.e., those in which Crippled Growth sectors occur. Right, the fact that the cytoplasmic and infectious factors responsible for Crippled Growth and Senescence amplify in different kinds of hyphae (only stationary ones for Crippled Growth when growing on M2 and apical ones for Senescence) is easily visualized by testing explants taken at various distances from the inoculation point. In the case of Crippled Growth, explants from the center of the culture regenerate sick cultures if inoculated on permissive medium, while explants from the outer part of the culture generate sick cultures in the case of Senescence</figcaption></figure>

Surprisingly, C accumulates during stationary phase even in conditions that did not permit the development of Crippled Growth sector such as growth on M2 (figure 73). An interesting property of Crippled Growth is its sensitivity to stresses. Indeed cultures growing Crippled can easily be forced to reverse to a Normal Growing regimen by stressing them. Intriguingly, all tested stresses are able to do so: heat shock, cold shock, presence of noxious substances or of osmolytes in large amounts, ultraviolet irradiation... One of the most efficient ones is physical stress. Indeed, shaking a Crippled Growing
mycelium with a Fastprep in order to break apart the hyphae results in fragments regenerating Normal
Growing-mycelia. Another way to revert Crippled Growth is to undergo a sexual cycle. As stated above

the Crippled Growing mycelium is female sterile. It is
however male fertile. Male gametes never transmit C.
Also, when sexual reproduction occurs, i.e., during
stationary phase C is present in maternal hyphae; yet
freshly germinated ascospores never present Crippled
Growth directly after germination. Overall, the
cytoplasmic and infectious factor C has a complex
determinism, which is summarized in figure 74:
induction by stationary phase, reversal by stresses
and role of growth condition, (i.e., development of
sectors only in medium containing yeast extract)

To understand the molecular mechanism
involved in the generation of C and the development
of Crippled Growth sectors, a classical genetic
approach has been undertaken. Note that purification

of C was not feasible because, in order to follow its presence during *in vitro* purification, it would have
been necessary to reintroduce it in hyphae, which could only be done by stressing them and thus
reverting Crippled Growth. Mutants impairing Crippled Growth (*IDC mutants*) and mutants promoting
Crippled Growth (*PDC mutants*) were selected (figure 71).

Two major classes of IDC mutants were retrieved. The first one still accumulated C during stationary phase and thus mutants from this class were likely constitutively-stressed during growth, which resulted in C elimination. Accordingly, all these mutants have additional phenotypes. The gene affected in only one of these mutants has been identified and encodes a potassium transporter. Defect in potassium transport can indeed be stressful during growth. The second class did not produce C, even in stationary phase and were thus likely affected in the pathway directly involved in the making of C. In addition to their effect on Crippled Growth, all these mutants have the same set of defects, including lack of pigments, aerial hyphae, anastomosis and development of fruiting bodies after fertilization. Interestingly, these are features occurring as the fungus enters into stationary phase, in accordance with

<figure><img src="image_79.png" /><figcaption>**Figure 74** Summary of the properties of C.</figcaption></figure>
<figure><img src="image_80.png" /><figcaption>**Figure 75** A model to account for Crippled Growth. See text for details.</figcaption></figure>

the induction of *C* during this phase. Identification of the genes mutated in the *IDC* and the *PDC1* mutants followed by reverse genetic analyses of candidate genes and phenotypic analyses of single and double mutants, led to the proposal of a model whereby the *C* element is a self-activated MAP kinase pathway (figure 75). On M2 medium, this pathway is normally off in growing hyphae thanks to an “Elimination Factor” and activates upon entrance into stationary phase. This is seen by the re-localization of the PaMpk1 MAP kinase inside nuclei specifically during stationary phase. Activation is prompted by the action of the PaNox1 NADPH oxidase complex, which produces ROS. The ROS signal likely travels through the IDC2 and IDC3 proteins that contain conserved cysteines crucial for their activity. It ends up in the PDC1 protein, which also contains conserved cysteines crucial for its activity. This protein represses the PaMpk1 pathway and is thus part of the “Elimination Factor”. Once lifted, activation of PaMpk1 can proceed and signal production of pigments, anastomosis, erection of hyphae and likely mobilization of
reserve to properly built the fruiting bodies. How activation is achieved is still mysterious since
phosphorylation of the TxY motif conserved in MAP kinases that usually activates these proteins is
constitutive. Activation is thus achieved through another mechanism as observed for some MAP kinases
in protists. Activation also requires an intact PaMpk2 MAP kinase pathway. Requirement of the PaMpk2
pathway for activity of PaMpk1 has been observed in related fungi. Upon growth renewal after transfer
on fresh medium, the “Elimination Factor” is reactivated, PaMpk1 is inactivated and growth can again
proceed normally.

On M2 + YE medium, the pathway is also usually off and hence growth proceeds normally. Like on M2, the pathway activates using the same actors during stationary phase. Nevertheless, on M2 + YE, the mechanism repressing C that contains PDC1 is inactive during growth. Thus upon transfer to fresh medium, PaMpk1 is not inactivated and Crippled Growth occurs because signaling for features specific of resting hyphae (pigment, anastomoses, erection of hyphae and preparation of sexual reproduction) are expressed in dividing ones . Similarly, during growth PaMpk1 can activate spontaneously and sectors of Crippled Growth ensue. This spontaneous activation of PaMpk1 may be effected by both the PaNox1 and PaNox2 NADPH oxidase complexes.

This model has been confirmed by many experiments, but much remains to be done to clearly
understand how C is created and how it affects growth. Many factors affecting Crippled Growth are
poorly characterized and their position in the pathway is unclear: IDC4, PaPsr1/PaWhi2, the STRIPAK
complex... Especially, the role of the transcription factor PaPro1 within the cascade is unclear. It has been
shown to activate the expression of some of the genes involved in Crippled Growth, but in a marginal
way. Whether this factor is also part of the self-activation of the pathway should to be determined.
Similarly, the genes in many IDC and especially in many PDC mutants remain to be identified. It is very
likely that the actual functioning of the PaMpk1 pathway during Normal and Crippled Growth is much
more complex than what the model of figure 75 depicts. Finally, we will further discuss this
PaMpk1/PaMpk2 pathway during the analysis of sexual reproduction and self-versus non-self-
recognition, since it is a major regulator of both phenomena.

In addition to Senescence and Crippled Growth, *P. anserina* and/or its sibling species is the
subject of additional phenotypic instabilities, i.e., repetitive variable appearance of the mycelium in well-
defined conditions. Foremost, all accumulate with great frequency *Incolore* mutations (figure 76). Indeed,
*Incolore* mutations are dominant and confer a selective advantage to nuclei that carry them: these divide