# The soil under the microscope: the optical examination of a small area of Scots pine litter (Pinus sylvestris L.)

Jean-François Ponge

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Jean-François Ponge. The soil under the microscope: the optical examination of a small area of Scots pine litter (Pinus sylvestris L.). 2010. hal-00488081

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# The soil under the microscope: the optical examination of a small area of Scots pine litter
(*Pinus sylvestris* L.)*

Jean-François Ponge

Muséum National d'Histoire Naturelle, CNRS UMR
7179, 4 avenue du Petit-Château, 91800 Brunoy,
France, E-mail: ponge@mnhn.fr

<footnote>* English translation of three papers published in French by Ponge (1984, 1985 and 1988). Some parts were rearranged for clarity but no attempt was made to update species and horizon names according to modern literature</footnote>
<table><tr><td>INTRODUCTION</td><td>9</td></tr><tr><td>Study site, sampling procedure and field observations</td><td>10</td></tr><tr><td>Preparation of the material for laboratory observation</td><td>11</td></tr><tr><td>THE L<sub>1</sub> LAYER: RECENTLY FALLEN NEEDLES AND LIVING MOSS</td><td>12</td></tr><tr><td>Decaying needles</td><td>15</td></tr><tr><td>Needles without any external sign of fungal attack</td><td>15</td></tr><tr><td>Needles attacked by parasitic fungi</td><td>17</td></tr><tr><td>Lophodermium pinastri</td><td>17</td></tr><tr><td>Ceuthospora pinastri and Lophodermella sp.</td><td>17</td></tr><tr><td>Needles attacked by the saprophytic Verticicladium trifidum</td><td>17</td></tr><tr><td>Needles attacked by Verticicladium trifidum after Lophodermium pinastri and Ceuthospora pinastri</td><td>19</td></tr><tr><td>Conclusions about decaying needles</td><td>21</td></tr><tr><td>Decaying twigs</td><td>21</td></tr><tr><td>Bark fragments</td><td>22</td></tr><tr><td>Living and decaying moss (Pseusceropodium purum)</td><td>23</td></tr><tr><td>Fauna</td><td>23</td></tr><tr><td>Inventory</td><td>23</td></tr><tr><td>Enchytraeid worms</td><td>23</td></tr><tr><td>Transport on the tegument</td><td>25</td></tr><tr><td>Feeding activity</td><td>27</td></tr><tr><td>Nematodes</td><td>27</td></tr><tr><td>Oribatid mites</td><td>27</td></tr><tr><td>Springtails</td><td>29</td></tr><tr><td>Fly larvae</td><td>29</td></tr><tr><td>Animal faeces</td><td>29</td></tr><tr><td>Animal corpses</td><td>31</td></tr><tr><td>Conclusions about the L<sub>1</sub> layer</td><td>31</td></tr></table>
<table>
  <tbody>
    <tr>
      <td>THE L<sub>2</sub> LAYER: DECAYING NEEDLES AND DEAD MOSS</td>
      <td>32</td>
    </tr>
    <tr>
      <td>Decaying pine needles</td>
      <td>34</td>
    </tr>
    <tr>
      <td>Needles attacked by Verticicladium trifidum</td>
      <td>35</td>
    </tr>
    <tr>
      <td>Needles attacked by Marasmius androsaceus</td>
      <td>35</td>
    </tr>
    <tr>
      <td>Needles attacked by unidentified fungi</td>
      <td>37</td>
    </tr>
    <tr>
      <td>Needles with bacterial development</td>
      <td>39</td>
    </tr>
    <tr>
      <td>Needles with bacterial and micro-algal development</td>
      <td>40</td>
    </tr>
    <tr>
      <td>Needles penetrated by enchytraeids</td>
      <td>39</td>
    </tr>
    <tr>
      <td>Hyphal networks embedding decaying pine needles</td>
      <td>41</td>
    </tr>
    <tr>
      <td>Conclusions about decaying pine needles</td>
      <td>44</td>
    </tr>
    <tr>
      <td>Decaying pine twigs</td>
      <td>43</td>
    </tr>
    <tr>
      <td>Decaying euphylls of pine</td>
      <td>45</td>
    </tr>
    <tr>
      <td>Decaying male flowers of pine</td>
      <td>45</td>
    </tr>
    <tr>
      <td>Decaying branchwood of pine</td>
      <td>45</td>
    </tr>
    <tr>
      <td>Bark</td>
      <td>46</td>
    </tr>
    <tr>
      <td>Wood</td>
      <td>47</td>
    </tr>
    <tr>
      <td>Pine ectomycorrhizae</td>
      <td>49</td>
    </tr>
    <tr>
      <td>Decaying moss (Pseudoscleropodium purum)</td>
      <td>49</td>
    </tr>
    <tr>
      <td>Stems</td>
      <td>49</td>
    </tr>
    <tr>
      <td>Leaves</td>
      <td>49</td>
    </tr>
    <tr>
      <td>Decaying bracken foliage (Pteridium aquilinum)</td>
      <td>50</td>
    </tr>
    <tr>
      <td>Leaflets</td>
      <td>50</td>
    </tr>
    <tr>
      <td>• Leaflets with limb still present</td>
      <td>50</td>
    </tr>
    <tr>
      <td>• Vein networks</td>
      <td>52</td>
    </tr>
    <tr>
      <td>Rachises</td>
      <td>52</td>
    </tr>
    <tr>
      <td>Fauna</td>
      <td>52</td>
    </tr>
    <tr>
      <td>Inventory</td>
      <td>53</td>
    </tr>
    <tr>
      <td>Enchytraeid worms</td>
      <td>56</td>
    </tr>
  </tbody>
</table>
<table>
  <tr>
    <td>Oribatid mites</td>
    <td>60</td>
  </tr>
  <tr>
    <td>Springtails</td>
    <td>62</td>
  </tr>
  <tr>
    <td>Fly larvae</td>
    <td>62</td>
  </tr>
  <tr>
    <td>Nematodes</td>
    <td>64</td>
  </tr>
  <tr>
    <td>Cyst amoebae</td>
    <td>64</td>
  </tr>
  <tr>
    <td>Animal faeces</td>
    <td>64</td>
  </tr>
  <tr>
    <td>Macroinvertebrate faeces</td>
    <td>64</td>
  </tr>
  <tr>
    <td>Enchytraeid faeces</td>
    <td>65</td>
  </tr>
  <tr>
    <td>Oribatid faeces</td>
    <td>65</td>
  </tr>
  <tr>
    <td>Springtail faeces</td>
    <td>69</td>
  </tr>
  <tr>
    <td>Animal corpses and corpse fragments</td>
    <td>69</td>
  </tr>
  <tr>
    <td>Conclusions about the L<sub>2</sub> layer</td>
    <td>71</td>
  </tr>
  <tr>
    <td>THE F<sub>1</sub> LAYER: FRAGMENTED NEEDLES, INVERTEBRATE FAECES AND MYCORRHIZAE</td>
    <td>73</td>
  </tr>
  <tr>
    <td>Decaying pine needles</td>
    <td>77</td>
  </tr>
  <tr>
    <td>Fungal mantle around pine needles</td>
    <td>77</td>
  </tr>
  <tr>
    <td>Inside of pine needles</td>
    <td>79</td>
  </tr>
  <tr>
    <td>Needles attacked by Verticicladium trifidum, but not by Cenococcum geophilum</td>
    <td>79</td>
  </tr>
  <tr>
    <td>Needles attacked by Cenococcum geophilum</td>
    <td>80</td>
  </tr>
  <tr>
    <td>Pine roots and mycorrhizae, mycorrhizal fungi</td>
    <td>82</td>
  </tr>
  <tr>
    <td>Living roots</td>
    <td>82</td>
  </tr>
  <tr>
    <td>Coralloid orange-brown ectomycorrhizae</td>
    <td>82</td>
  </tr>
  <tr>
    <td>Black ectomycorrhizae</td>
    <td>83</td>
  </tr>
  <tr>
    <td>Pink ectomycorrhizae</td>
    <td>84</td>
  </tr>
  <tr>
    <td>Long roots</td>
    <td>84</td>
  </tr>
  <tr>
    <td>Decaying roots</td>
    <td>86</td>
  </tr>
  <tr>
    <td>Short roots</td>
    <td>86</td>
  </tr>
  <tr>
    <td>Long roots</td>
    <td>87</td>
  </tr>
</table>
<table>
  <tbody>
    <tr>
      <td>• Sclerotia of Cenococcum geophilum</td>
      <td>87</td>
    </tr>
    <tr>
      <td>Pine twigs</td>
      <td>87</td>
    </tr>
    <tr>
      <td>Preliminary remarks</td>
      <td>87</td>
    </tr>
    <tr>
      <td>Wood</td>
      <td>88</td>
    </tr>
    <tr>
      <td>• Penetration by basidiomycetes without any attack of lignified walls</td>
      <td>88</td>
    </tr>
    <tr>
      <td>• Penetration by fungi and bacteria with soft-rot attack</td>
      <td>88</td>
    </tr>
    <tr>
      <td>Bark</td>
      <td>90</td>
    </tr>
    <tr>
      <td>Pine bark</td>
      <td>90</td>
    </tr>
    <tr>
      <td>Notice</td>
      <td>90</td>
    </tr>
    <tr>
      <td>Thin bark fragments</td>
      <td>90</td>
    </tr>
    <tr>
      <td>Thick bark fragments</td>
      <td>91</td>
    </tr>
    <tr>
      <td>Pine euphylls</td>
      <td>91</td>
    </tr>
    <tr>
      <td>Male flowers of pine</td>
      <td>92</td>
    </tr>
    <tr>
      <td>Miscellaneous pine material</td>
      <td>92</td>
    </tr>
    <tr>
      <td>Leafy shoots of moss (Pseudoscleropodium purum)</td>
      <td>93</td>
    </tr>
    <tr>
      <td>Preliminary remarks</td>
      <td>93</td>
    </tr>
    <tr>
      <td>Leaves</td>
      <td>93</td>
    </tr>
    <tr>
      <td>• Attack of soft-rot type</td>
      <td>93</td>
    </tr>
    <tr>
      <td>• Attack of white-rot type</td>
      <td>93</td>
    </tr>
    <tr>
      <td>Stems</td>
      <td>93</td>
    </tr>
    <tr>
      <td>• Notice</td>
      <td>95</td>
    </tr>
    <tr>
      <td>• Cells with cell contents still present</td>
      <td>95</td>
    </tr>
    <tr>
      <td>• Penetration of fungal hyphae</td>
      <td>95</td>
    </tr>
    <tr>
      <td>• Penetration of microfauna</td>
      <td>95</td>
    </tr>
    <tr>
      <td>• Hollow moss stems</td>
      <td>96</td>
    </tr>
    <tr>
      <td>• Concluding remarks</td>
      <td>96</td>
    </tr>
    <tr>
      <td>Bracken fronds (Pteridium aquilinum)</td>
      <td>96</td>
    </tr>
    <tr>
      <td>Bracken leaflets</td>
      <td>96</td>
    </tr>
  </tbody>
</table>
<table>
  <tbody>
    <tr>
      <td>Bacterial stage</td>
      <td>98</td>
    </tr>
    <tr>
      <td>Cenococcum stage</td>
      <td>98</td>
    </tr>
    <tr>
      <td>Rachises</td>
      <td>98</td>
    </tr>
    <tr>
      <td>Birch (Betula pubescens Ehrh.)</td>
      <td>99</td>
    </tr>
    <tr>
      <td>Leaves</td>
      <td>99</td>
    </tr>
    <tr>
      <td>Achenes</td>
      <td>99</td>
    </tr>
    <tr>
      <td>Mucus</td>
      <td>101</td>
    </tr>
    <tr>
      <td>Preliminary remarks</td>
      <td>101</td>
    </tr>
    <tr>
      <td>Fungal development</td>
      <td>101</td>
    </tr>
    <tr>
      <td>Decay of the fungus</td>
      <td>101</td>
    </tr>
    <tr>
      <td>Rhizomorphs</td>
      <td>101</td>
    </tr>
    <tr>
      <td>Quartz grains</td>
      <td>103</td>
    </tr>
    <tr>
      <td>Spider cocoons</td>
      <td>103</td>
    </tr>
    <tr>
      <td>Colonies of Chrysophyceae (golden algae)</td>
      <td>103</td>
    </tr>
    <tr>
      <td>Fauna</td>
      <td>103</td>
    </tr>
    <tr>
      <td>Inventory</td>
      <td>103</td>
    </tr>
    <tr>
      <td>Protozoa</td>
      <td>105</td>
    </tr>
    <tr>
      <td>Rotifers</td>
      <td>107</td>
    </tr>
    <tr>
      <td>Nematodes</td>
      <td>107</td>
    </tr>
    <tr>
      <td>Enchytraeid worms</td>
      <td>108</td>
    </tr>
    <tr>
      <td>Preliminary remarks</td>
      <td>108</td>
    </tr>
    <tr>
      <td>Generalities</td>
      <td>108</td>
    </tr>
    <tr>
      <td>Moss material</td>
      <td>109</td>
    </tr>
    <tr>
      <td>Pine tissues</td>
      <td>109</td>
    </tr>
    <tr>
      <td>Fungal material</td>
      <td>109</td>
    </tr>
    <tr>
      <td>Bacteria</td>
      <td>110</td>
    </tr>
    <tr>
      <td>Thecamoeban cysts</td>
      <td>110</td>
    </tr>
    <tr>
      <td>Pine pollen</td>
      <td>110</td>
    </tr>
  </tbody>
</table>
<table>
  <tbody>
    <tr>
      <td>• Mesofaunal faeces</td>
      <td>110</td>
    </tr>
    <tr>
      <td>• Algae</td>
      <td>112</td>
    </tr>
    <tr>
      <td>• Intestinal microflora</td>
      <td>112</td>
    </tr>
    <tr>
      <td>• Transport on the tegument</td>
      <td>112</td>
    </tr>
    <tr>
      <td>Oribatid mites</td>
      <td>112</td>
    </tr>
    <tr>
      <td>• Generalities</td>
      <td>112</td>
    </tr>
    <tr>
      <td>• Box mites</td>
      <td>114</td>
    </tr>
    <tr>
      <td>Nothrus sylvestris</td>
      <td>115</td>
    </tr>
    <tr>
      <td>• Oppiidae</td>
      <td>115</td>
    </tr>
    <tr>
      <td>• Other species</td>
      <td>116</td>
    </tr>
    <tr>
      <td>• Conclusions</td>
      <td>116</td>
    </tr>
    <tr>
      <td>Miscellaneous mites and spiders</td>
      <td>116</td>
    </tr>
    <tr>
      <td>Woodlice</td>
      <td>117</td>
    </tr>
    <tr>
      <td>Springtails</td>
      <td>117</td>
    </tr>
    <tr>
      <td>Insect larvae</td>
      <td>120</td>
    </tr>
    <tr>
      <td>Animal corpses</td>
      <td>121</td>
    </tr>
    <tr>
      <td>Preliminary remarks</td>
      <td>121</td>
    </tr>
    <tr>
      <td>Enchytraeid worms</td>
      <td>122</td>
    </tr>
    <tr>
      <td>Oribatid mites</td>
      <td>122</td>
    </tr>
    <tr>
      <td>Other arthropods</td>
      <td>125</td>
    </tr>
    <tr>
      <td>Animal faeces</td>
      <td>125</td>
    </tr>
    <tr>
      <td>Type A faeces (slugs?)</td>
      <td>125</td>
    </tr>
    <tr>
      <td>Type B faeces (woodlice?)</td>
      <td>126</td>
    </tr>
    <tr>
      <td>Type C faeces (earthworms)</td>
      <td>127</td>
    </tr>
    <tr>
      <td>Mineral faeces</td>
      <td>129</td>
    </tr>
    <tr>
      <td>• Preliminary remarks</td>
      <td>129</td>
    </tr>
    <tr>
      <td>• White mineral faeces</td>
      <td>129</td>
    </tr>
  </tbody>
</table>
<table><tr><td>• Grey mineral faeces</td><td>130</td></tr><tr><td>Enchytraeid faeces</td><td>130</td></tr><tr><td>Oribatid faeces</td><td>130</td></tr><tr><td>Sciarid faeces</td><td>132</td></tr><tr><td>Conclusions about the F<sub>1</sub> layer</td><td>132</td></tr><tr><td>ACKNOWLEDGEMENTS</td><td>134</td></tr><tr><td>REFERENCES</td><td>135</td></tr></table>
INTRODUCTION

The purpose of the present document is to show the interest of morphological studies for a better understanding of the biological interactions which take place in the soil ecosystem. Soil micromorphology was mainly developed following studies by Jongerius (1957) and Babel (1964), but Hesselmann's (1926) microstratification and observations and methodologies by Kubiëna (1938) set the stage for a science which associates the knowledge of the mineral world to that of the living world. On this base Bal (1970) and thereafter Brun (1978) were able to perform integrated studies which confirmed the interest of observing the soil for understanding its functioning as a living system.

On their part, zoologists made observations which allowed them to demonstrate that earthworms were not the only group playing a driving role in the building of soil structure (Zachariae, 1964; Van der Drift, 1964). Accordingly, the role of bacterial colonies in the formation of soil structure, in particular of soil aggregates, was demonstrated by microbiologists (Proth, 1978). This decisive entrance of biology in the field of soil science allowed the scientific community to realize how it was difficult to achieve an integrated study at a fine level of resolution.

Heterogeneity is great in the soil ecosystem and results obtained in a given micro-site cannot be transposed to another. In order to reach representativeness over a given site or a given soil type, the specialist must multiply laboratory experiments, observations and measurements, being forced to become specialized on particular taxonomic groups. In every case, the specialist is able to demonstrate the paramount importance of the group he/she is studying, to the detriment of a comprehensive understanding of soil biological functioning. However, several studies showed that biological interactions play a prominent role in the relationships between microbes, animals and the mineral substrate (Kilbertus & Vannier, 1979; Parkinson et al., 1979; Touchot et al., 1983).

In opposition to the multiplication of observations, measurements and experiments, it can be scheduled to take into account the widest possible array of interactions taking place in a restricted volume of soil. If we consider that most typical processes taking place in the soil ecosystem, such as plant litter decomposition, involve more vertical than horizontal movements of matter, then studying a profile, even on a restricted area, should provide a coherent view of soil biological functioning at this right place. For that purpose each layer or horizon will be considered as a source of information on the past history of the just overlying layer or horizon, using traces of biological activity in it as clues. This postulate is unavoidable in such an integrated study. It should be noted that the local examination of litter and soil must be replaced in the context of forest soil heterogeneity. My purpose is not to ascribe my results to a site, to a soil or to a humus type, but just to study a microcosm where interactions between soil organisms (microbes and animals), plant organic matter (litter and root system) and mineral particles (sand, silt and clay) could be studied in the most comprehensive manner within the limits of sampling and observational techniques.

In the following text, the term 'humus' will designate every part of the soil with a high amount of organic matter, i.e. for the soil under consideration the organic layers L, F, H, and the A₁ horizon, and not any type of organic matter which could be isolated by physical or chemical agencies. Bal (1970) underwent a sound discussion of the meaning of 'humus'.
Study site, sampling procedure and field observations

The humus profile under study is a dysmoder (sensu Manil, 1959) under Scots pine (*Pinus sylvestris* L.). The forest stand is a 35-yr-old plantation located in the Forêt d'Orléans (Loiret, France). The study site has been studied by Poursin & Ponge (1984) and Arpin et al. (1986a, b), which gave more details about it. The soil is leached acidic sandy with temporary waterlogging and is covered by a continuous carpet of the moss *Pseudoscleropodium purum* (L.), known for its common association with coniferous stands (Bournéias, 1979). Fronds of bracken, *Pteridium aquilinum* (L.), are sparsely distributed in the understory. The present paper deals only with the topmost three layers where most biological activity was shown to take place.

Sampling took place in August 1981. The different layers and soil horizons were separated directly in the field over a 5 x 5 cm area, using fingers and fine scissors, and were immediately fixed and preserved in ethyl alcohol (95% v/v) in separate plastic jars until study. The method of identification of litter layers and soil horizons by Babel (1971) was used, but we changed the nomenclature (Table 1). Babel's sub-horizons were called layers, considering that (i) organic deposits should not be confused with soil horizons, according to the opinion by Bal (1970), and (ii) faunistic and microbiological differences are such between sub-layers that they should be erected to the layer rank.

Table 1 summarizes most observations which were done on the field with the naked eye and were used for separating organic layers and first organo-mineral horizons of the topsoil. The top two layers are made of a continuous carpet of *Pseudoscleropodium purum*, living in L₁ (Fig. 1), dead in L₂ (Fig. 2). Thus the thickness of L₁ and L₂ does not correspond to freely accumulating pine litter: in these two layers most plant material was made of leafy shoots of mosses.

<table><caption>**Table 1.** Microstratification: qualitative features of the different layers and horizons (+++ = dominant in the layer/horizon considered; ++ = abundant but not dominant; + = present but not abundant)</caption><thead><tr><th>Layers Horizons</th><th>Thickness (cm)</th><th>Transition with the underlying</th><th>Moss</th><th>Needles</th><th>Mycelia</th><th>Mycorrhizae</th><th>Faeces</th><th>Amorphous organic matter</th><th>Sand</th><th>Iron spots</th></tr></thead><tbody><tr><td>L<sub>1</sub></td><td>3</td><td>Fuzzy</td><td>+++ living</td><td>+ brown entire</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>L<sub>2</sub></td><td>2</td><td>Sharp</td><td>++ dead</td><td>++ Black entire</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>F<sub>1</sub></td><td>1</td><td>Fuzzy</td><td></td><td>+++ fragments</td><td>++</td><td></td><td>+</td><td></td><td></td><td></td></tr><tr><td>F<sub>2</sub></td><td>1</td><td>Fuzzy</td><td></td><td>++ fragments</td><td></td><td>+</td><td>++</td><td></td><td></td><td></td></tr><tr><td>H<sub>1</sub></td><td>0.5</td><td>Sharp</td><td></td><td>+ fragments</td><td></td><td>+</td><td>+++</td><td></td><td></td><td></td></tr><tr><td>H<sub>2</sub></td><td>0.5</td><td>Fuzzy</td><td></td><td></td><td></td><td></td><td>+</td><td>+++</td><td>++</td><td></td></tr><tr><td>A<sub>11</sub></td><td>1 to 4</td><td>Sharp</td><td></td><td></td><td></td><td></td><td></td><td>++</td><td>++</td><td></td></tr><tr><td>A<sub>12</sub></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td>+++</td><td>+</td></tr></tbody></table>

Winter-spring waterlogging was visible through the presence of rusty spots in A₁₂, due to seasonal anoxia, and the transformation of animal faeces into an amorphous mass in H₂ (partly incorporated in A₁₁, with a sharp transition to A₁₂), due to groundwater movements. These features are
typical of hydromorphic forms of moder or hydromoder (Duchaufour, 1977). Measurements of pH<sub>water</sub> (Poursin & Ponge, 1984) indicated a value of 3.3, which is very acid. This, together with the total volume of accumulated organic matter in so a young forest stand and the sharp transition between A<sub>11</sub> and A<sub>12</sub>, prompted me to interpret this humus profile as transitional to mor at the time of the study. In the A<sub>12</sub> horizon, discoloured and spotted with rusty spots, there were rhizomes of *Pteridium aquilinum* and woody roots of *P. sylvestris*, some of which were decayed and transformed in dark masses of animal faeces, with a sharp transition to the surrounding bleached sandy horizon.

## Preparation of the material for laboratory observation

Early observations were made on litter/humus material after spreading it in petri dishes filled with alcohol. These preliminary observations (accompanied or not by counts when necessary) were done under a dissecting microscope at varying magnification (x 10 to x 40). After a thorough examination of the whole material, specimens of the variety of decay stages (or other cause of variation) in the various plant organs were observed with a light microscope at higher magnification (x 100 to x 400), with or without phase contrast. Plant organs of varying nature (needles, twigs, bark fragments, roots, moss and bracken leaves and stems) were sectioned with a microtome (7.5 µm thickness) after dehydration in absolute ethyl alcohol and embedding in paraffin then sections were mounted either in chloral-lactophenol (to be observed under phase contrast) or in methylblue-Sudan III-lactophenol (to be observed in natural light). Methylblue stains indiscriminately cell organelles, allowing us to distinguish living bacterial and fungal cells within plant tissues, while Sudan III stains in red waxes, cutin and suber, and at last phase contrast was useful to study dead cell parts at varying stages of transformation (Locquin & Langeron, 1978). Following passage from ethyl alcohol to toluene, resins were badly conserved in sectioned plant material. Animal faeces were observed from the outside under a binocular microscope then mounted directly in lactophenol after squashing them or part of them under coverslides.

All animals, whatever their size (nematodes were collected as well as bigger fauna), were collected and identified at the species level whenever possible. Poorly sclerotized animals such as enchytraeids (potworms), springtails, some mites, insect larvae and nematodes were observed by transparency under phase contrast after mounting them in chloral-lactophenol, for the sake of identification and in order to examine their gut contents (when solid particles had been ingested). Heavily sclerotized animals (oribatid mites) were dissected then their gut parts were mounted in the same manner as above. The biovolume of each mounted animal was calculated using an approximate method taking into account the fact the actual volume of the body was lesser than that of a parallelepiped of same length, width and thickness, and higher than that of an octahedron of same dimensions, the ratio between both values being 4:1. It can be easily demonstrated that this approximation is true whatever the size and the shape of the animal, provided length, width and thickness could be calculated, and the animal was in a good state of turgor (which was insured by mounting it in chloral-lactophenol). The more an animal was squashed on the microscopic slide, the more a parallelepiped approximated its true biovolume. Corpses (empty cuticles of arthropods) were not measured because their volume could not be approximated with the above method.
THE L₁ LAYER: RECENTLY FALLEN NEEDLES AND LIVING MOSS

<figure><img src="image_2.png" /><figcaption>**Figure 1.** The L₁ layer before fixation and transport to the laboratory</figcaption></figure>

A volume of ca. 50 mL of L₁ layer was sorted under the binocular microscope, which allowed me to separate different components, the volume of which was measured by meniscus displacement in graduated columns (Table 2). Exception was for animals, the biovolume of which was measured individually at higher magnification, as explained above. Animal faeces and fungal mycelia (mostly basidiomycetes), which were poorly represented and firmly appressed to their plant support (moss shoots, pine needles), were included in plant components.

<table><caption>**Table 2.** Volume of the different components of the L₁ layer sorted under the binocular microscope</caption><thead><tr><td>Litter components</td><td>Volume (µL)</td></tr></thead><tbody><tr><td>Moss</td><td>7500 ± 100</td></tr><tr><td>Pine needles</td><td>1000 ± 100</td></tr><tr><td>Pine bark</td><td>900 ± 100</td></tr><tr><td>Pine twigs</td><td>300 ± 100</td></tr><tr><td>Male flowers of pine</td><td>18 ± 2</td></tr><tr><td>Living fauna</td><td>0.4 – 2</td></tr><tr><td>Total</td><td>9700 ± 100</td></tr></tbody></table>

Table 2 shows that the total volume of litter components (~ 10 mL) was much less than the volume of layer sampled (50 mL), thereby indicating the importance of voids in this well-aerated environment.
The biovolume of fauna seems very weak compared to that of the plant material. However, it must be noticed that this superficial layer was the seat of the movement of big invertebrates which lived and moved on the ground surface, but could be temporary present or not. At the time and at the right place of sampling no woodlouse, millipede, centipede or lumbricid worm was observed, which would have been magnified the biovolume of fauna to a great extent, without increasing its role. This random factor will be neglected in the course of my study by taking into account mainly traces of animal activity, onto which my estimates of the respective impact of different animal groups will be based. However, living animals which were sorted under the binocular microscope allowed me (i) to study the transformation of plant and microbial substrates during intestinal transit time, and (ii) to assign faeces to animal groups. In the L₁ layer, I observed only a weak number of faeces, all of them were attributed to oribatid mites.

Sorting litter components under the binocular microscope allowed me to show the dominance of living shoots of the moss *Pseudoscleropodium purum* and the visible development of the ascomycete *Lophodermium pinastri* (Schrad.) Chev., which was reported in nearly all recently fallen pine needles (Fig. 1). This fungus formed black buttonhole-like subcuticular ascocarps (Ø 1 mm) and diaphragms which were visible from the outside and allowed identifying it with certitude (Minter et al., 1978; Minter, 1981). Figures 2 and 3 show parts of pine needles which were colonised by this fungus, known to infest foliage falling in spring or summer (Jones, 1935; Lanier, 1968).

Dating of the plant material present in the L₁ layer was possible thanks to the presence of male flowers of pine (know to fall at the end of June or beginning of July) and to the absence of euphylls (bracts, present in inflorescences and shoots of the year, known to fall at the beginning of June): I could conclude that the material present in L1 had fallen during July and beginning of August.

Some needle parts appeared to be colonised by two other ascomycetes. *Ceuthospora pinastri* (Fr.) Höhn.* is the imperfect stage of *Phacidium lacerum* Fr. (Gremmen, 1959). I did not observe its pycnidia in litterfall baskets which were installed in the same site, but Hayes (1965) indicated that it fructifies only once the needle has fallen on the ground. Most needles infested by *Ceuthospora pinastri* showed signs of previous attack by *Lophodermium pinastri*, the former colonizing zones which had not been colonised by the latter (Fig. 3). *Ceuthospora pinastri* appeared in the form of numerous black subhypodermic pycnidia (Fig. 4), often grouped by two or three. *Lophodermella* von Höhn., which was not identified to species level by lack of mature fructifications, formed sub-hypodermic ascocarps (Fig. 5), contrary to *Lophodermium*, ascocarps of which are subepidermal or subcuticular according to species. *Lophodermella* is known to cause precocious fall of pine needles (Mitchell et al., 1978b) and this was verified through the small size of pine needles which were infected by this fungus.

Needles which were in contact with the underlying L₂ layer (the passage from L₁ to L₂ was progressive, see Table 1) were colonised by *Verticicladium trifidum* Preuss, the imperfect stage of the ascomycete *Desmazierella acicola* Lib. This fungus was easily recognized by its black erected conidiophores (Fig. 6), protruding through stomatal apertures from melanised stromata located in substomatic chambers. At an early stage, only substomatic chambers and ostioles appear in black, corresponding to the 'black stomata' stage of Hayes (1965). Thereafter, as far as the hypodermis becomes melanised, needles turn to black on their whole surface and become covered with a carpet of conidiophores of this fungus. *Verticicladium trifidum* was recognized as a major secondary coloniser

<footnote>* The identification of this fungus was confirmed by Dr B.C. Sutton (Commonwealth Mycological Institute, Kew, Surrey, UK) who is warmly acknowledged</footnote>
<figure><img src="image_3.png" /><figcaption>Fig. 2: Needle infested by *Lophodermium pinastri*; Fig. 3: Needle infested by *Lophodermium pinastri* and *Ceuthospora pinastri*; Fig. 4: Subhypodermic pycnidium of *Ceuthospora pinastri* (BMSL); Fig. 5: Subhypodermic immature ascocarp of *Lophodermella* (CP); Fig. 6: Conidiophores of *Verticicladium trifidum* (CP); Fig. 7: Pine needle attacked by fauna; Fig. 8: Idem, other mode of attack; Fig. 9: Light-brown pine needle without any fungal fructification, section through epidermis, hypodermis and mesophyll at the level of a stomatal aperture (BMSL)</figcaption></figure>

CP = phase contrast; BMSL = methyl blue-Sudan III-lactophenol coloration; d. = diaphragms; a. = ascocarps; c. = cavities caused by animal biting; tun. = faunal tunnels; ep. = epidermis; hyp. = hypodermis; L.p. = *Lophodermium pinastri*; C.p. = *Ceuthospora pinastri*; length of segment = 50 µm, unless otherwise indicated
of Scots pine needles by Kendrick & Burges (1962). However, in my sample, I also observed it as a
primary coloniser.

At the level of the L₁ layer, the impact of soil animals was but feeble. Three kinds of traces of faunal attack were observed on pine needles, (i) small disjoint cavities, aligned along stomatal lines (Fig. 8), maybe indicating that substomatic stromata of *Verticicladium trifidum* were consumed (?), (ii) superficial grooves (Fig. 7), or (iii) tunnels at the inside of needles (not visible from the outside, see sections shown in Figs. 24 & 25). All these attacks were attributed to enchytraeids, for reasons which will be enlightened during the study of the underlying layer L₂ where these animals were abundant at the time of sampling. However, it must be noticed that such features of biting or tunnelling were exceptional in L₁, faunal feeding activity being mainly devoted to browsing of conidiophores and superficial mycelia, as was shown by the examination of gut contents.

Decaying needles

All needle types were sectioned (15 different morphological types), but only four of them will be
presented here, as being particularly demonstrative.

Needles without any external sign of fungal attack

Some green needles were found in the L₁ layer. Sections revealed that they were identical to living foliage: chloroplasts (stained violet) were intact and no sign of fungal infestation was visible. These needles, which fell precociously probably because of branch fall or strong winds, are only mentioned for the record.

Some needles, although light-brown-coloured, did not show any visible trace of fungal
development, to the exception of some hyaline hyphae crawling at their surface. When sectioned, these
needles did not appear to be intact. They were invaded at the inside by a network of hyaline hyphae,
without any fructification. Only living cells of pine seemed to be attacked, i.e. mesophyll and
endoderm cells, living cells of transfusion tissue, phloem and vascular rays. In these tissues, cell walls
were brown-coloured (Figs. 9 & 10), indicating a biochemical transformation of cellulose. Lignified
dead cells (sclerenchyma, metaxylem tracheids with bordered pits, transfusion tracheids) were not
invaded and lignin remained transparent (Fig. 11). Living cells with lignified walls (epidermis) were
not invaded (Fig. 9). In all cases cytoplasm was replaced by hyphae of this fungus and cellulose
seemed to be modified. Lignin seemed to arrest the development of the fungus, which was not able to
penetrate tracheids through the areolae of bordered pits. However, suberin did not impede thin cell
walls of the endodermis to be penetrated.

In the mesophyll, hyphae where seen growing in every part of the cells (Figs. 9 & 10). I
hypothesize that this fungus was able to destroy cellulosic walls once cytoplasmic contents have been
digested.

Penetration of the needles took place through stomatal apertures, given the dense development
<figure><img src="image_4.png" /><figcaption>Fig. 10: Light-brown pine needle without any fungal fructification, transversal section through the mesophyll (BMSL); Fig. 11: Idem, transversal section through phloem, vascular cambium, metaxylem and protoxylem (BMSL); Fig. 12: Idem, transversal section showing hyphae appressed to the cuticle (BMSL); Fig. 13: Idem, hyphae penetrating through a stomatal aperture (BMSL); Fig. 14: Needle attacked by *Lophodermium pinastri*, longitudinal section showing hyphae in the mesophyll (CP); Fig. 15: Idem, transversal section showing transfusion tissue, xylem, phloem, perivascular sclerenchyma (BMSL); Fig. 16: Idem, longitudinal section showing melanised hyphae penetrating through a stomatal aperture and forming a melanised stroma (CP); Fig. 17: Needle attacked by *Verticicladium trifidum*, longitudinal section through phloem (BMSL)</figcaption></figure>

CP = phase contrast; BMSL = methyl blue-Sudan III-lactophenol coloration; h.h. = hyaline hyphae; h.m. = melanised hyphae; p.c. = cell walls; s.m. = melanised stromata; length of segment = 50 µm
of hyphae which was observed in sub-stomatal chambers (Fig. 9) and the network of hyphae of similar size and shape which was observed externally (Fig. 12). Penetration through stomata was also occasionally observed (Fig. 13).

Light-brown needles remained turgescent. Not any identity could be given to this fungus, since sporulation did not occur in my sample, and the parasitic or saprophytic nature of the infestation could not be revealed to the observer. However, it must be noticed that early colonisation of pine needles by fungi seemed to be the rule in the studied sample, even when no external sign of fungal attack was visible.

*Needles attacked by parasitic fungi*

* *Lophodermium pinastri*

A network of invading hyphae was clearly visible in the mesophyll (Fig. 14). Hyphae were thin-walled and segmented, with frequent bulb-like expansions (haustoria?). However, they were never seen penetrating mesophyll cells, which were collapsed and brown-coloured (more than with the former, unidentified fungus). Phloem appeared also brown and collapsed, but xylem tracheids and sclerenchyma cells remained transparent (Fig. 15). Thus lignin, contrary to cellulose, did not seem to be attacked, at least at this stage.

Melanised stromata were often observed, stemming from hyphae crawling at the surface of needles which further penetrated the needles through stomatal apertures (Fig. 16). These stromata were interpreted as resting stages of *Verticicladium trifidum*, awaiting senescence of internal colonies of *Lophodermium pinastri*. This might explain the observed succession between *Lophodermium pinastri* and *Verticicladium trifidum* observed on ageing needles.

* *Ceuthospora pinastri* and *Lophodermella sp.*

Needles attacked by these two fungi exhibited the same signs than when attacked by *Lophodermium pinastri*, i.e. colonisation of the mesophyll by a network of extra-cellular hyphae, with concomitant collapse and browning of cell walls, without any degradation of lignin.

*Needles attacked by the saprophytic Verticicladium trifidum*

The internal development of this ascomycete (conidial stage of *Desmazierella acicola*) was characterised by long thick-walled hyaline hyphae stretched along cell files of phloem (Fig. 17), protoxylem (Fig. 18) and resin ducts (Fig. 19). Mesophyll cells were also invaded as well as those of
<figure><img src="image_5.png" /><figcaption>Fig. 18: Needle attacked by *Verticicladium trifidum*, longitudinal section through protoxylem and metaxylem (BMSL); Fig. 19: Idem, transversal section of a needle showing epidermis, hypodermis, resin duct and thick-walled hyaline hyphae (BMSL); Fig. 20: Idem, transversal section through transfusion tissue showing thick-walled hyaline hyphae appressed to intact pine cell walls (BMSL); Fig. 21: Idem, tangential section through hypodermis showing guard cells invaded by hyaline hyphae and their walls impregnated by dark deposits (BMSL); Fig. 22: Idem, longitudinal section through a stomatal guard cell showing hyaline hyphae at the inside (BMSL); Fig. 23: Needle attacked by *Lophodermium pinastri* and *Ceuthospora pinastri* then *Verticicladium trifidum* and penetrated by an enchytraeid, longitudinal section showing cuticle covered with melanised hyphae, epidermis, hypodermis with black deposits, mesophyll with bacteria (BMSL); Fig. 24: Idem, longitudinal section showing the passage from a zone attacked by fungi only (mesophyll cell files still present) to a zone tunnelled by an enchytraeid, with faeces deposition (BMSL); Fig. 25: Idem, longitudinal section showing enchytraeid faces accumulated under the hypodermis, incrusted by black deposits (BMSL)</figcaption></figure>

BMSL = methyl blue-Sudan III-lactophenol coloration; h.h. = hyaline hyphae; h.m. = melanised hyphae; s.m. = melanised stromata; i.n. = black deposits; t.m. = mesophyll cell files; d.e. = enchytraeid faeces; length of segment = 50 µm
transfusion tissue (Fig. 20), into which starch grains could still be observed in abundance. The latter
observation was indirect evidence that the needle was functional when still on the tree, contrary to
needles previously parasitized by *Lophodermium pinastri*, *Ceuthospora pinastri* or *Lophodermella sp.*
*Verticicladium trifidum*, known as a soil saprophyte (Gremmen, 1977), can be considered in the
present case as a primary coloniser of fallen pine needles. At this stage, mesophyll, phloem,
protoxylem and transfusion tissue were invaded, and only metaxylem was still untouched. Cell walls
did not appear transformed, only cytoplasm being probably digested.

Melanised stromata of *Verticicladium trifidum* were observed in sub-stomatal chambers and stomatal apertures (Fig. 22). Melanin or melanin-like black substances seemed to diffuse and impregnate the walls of stomatal guard cells (Fig. 21) and hypodermis at the contact of melanised stromata, while hyaline hyphae invaded the inside of guard cells (Fig. 22).

The pine needles invaded by this fungus were brown-coloured, with lines of black stomata. At this stage conidiophores of *Verticicladium trifidum* were still not observed protruding singly or grouped by two or three from melanised stromata filling stomatal apertures.

*Needles attacked by Verticicladium trifidum after Lophodermium pinastri and Ceuthospora pinastri*

Like when *Verticicladium trifidum* was a primary coloniser, the presence of this fungus could be
attested by its thick long hyphae colonizing longitudinally the needles and by its melanised stromata
filling stomatal apertures. However, in the present case, blackening was generalized through the
hypodermis, forming a thin opaque layer which gave a black aspect to the whole needle (Fig. 23).
Previous colonisation by *Lophodermium pinastri* and *Ceuthospora pinastri* was attested by the
presence of melanised diaphragms of the former (Fig. 23) and subhypodermic pycnidia of the latter.

The studied needle showed traces of tunnelling by an animal which deposited its faeces at the
inside of the cavity it created. Figure 24 shows in the mesophyll the passage from a zone attacked by
fungi only (mesophyll cell files still recognizable) to a zone which had been ingested then transformed
into faeces by an animal. These faeces were round masses ca. 50-70 µm diameter closely packed under
the blackened hypodermis, where they replaced the mesophyll (Fig. 25). Stromatic diaphragms of
*Lophodermium pinastri* were not consumed, probably because of their toughness, and formed barriers
to the tunnelling activity of the animal: the upper right side of Figure 25 also shows such a black
diaphragm, together with an ascocarp which had been emptied after spore discharge (cuticle was
peeled off). In other places vascular and transfusion tissues could be attacked as well.

Through observations which were made in L₁, L₂ and F₁ layers, the tunnelling activity scarcely
observed in L₁ was attributed to enchytraeids. At the inside of enchytraeid faeces plant cells were no
longer recognizable, to the exception of some particular anatomical features. Cellulosic cell walls were
crushed and lacked transparency (Fig. 26), but lignified walls of tracheids were still intact and
recognizable (Fig. 27), indicating that vascular tissues were consumed. Hyaline hyphae (of unknown
origin) were observed but they were dead (not stainable). Bacterial development was noticed
everywhere, in some parts in the form of actively growing colonies (Fig. 28).
<figure><img src="image_6.png" /><figcaption>Fig. 26: Needle attacked by *Lophodermium pinastri* and *Ceuthospora pinastri* then *Verticicladium trifidum* and penetrated by an enchytraeid, detail of enchytraeid faeces showing plant cell wall remains and bacteria (CP); Fig. 27: Idem, detail of enchytraeid faeces showing remains of tracheids with bordered pits (metaxylem) (CP); Fig. 28: Idem, detail of enchytraeid faeces showing bacterial colonies (CP); Fig. 29: Idem, detail of basidiomycetous mycelium embedding the needle (CP); Fig. 30: Two-yr-old pine twig, transversal section showing suber, hypodermis and epidermis lifted off by a fungal pycnidium (BMSL); Fig. 31: Idem, transversal section showing medullar cells with intracellular hyphae (BMSL); Fig. 32: Idem, transversal section showing cortical cells, collapsed and with brown walls (BMSL); Fig. 33: Idem, transversal section showing metaxylem, medullar ray and vascular ray (BMSL)</figcaption></figure>

CP = phase contrast; BMSL = methyl blue-Sudan III-lactophenol coloration; a.r. = bordered pits; c.b. = bacterial colonies; b.m. = clamp connections; h.h. = hyaline hyphae; r.v. = vascular rays; r.m. = medullary rays; B.E. = earlywood; B.H. = latewood; length of segment = 50 µm
## Conclusions about decaying needles

Decay processes observed in L₁ pine needles were mainly due to the activity of pine-specific ascomycetes. Fungal development was mainly internal (to the exception of conidiophores of *Verticicladium trifidum*) but fungal hyphae were seen at the surface of needles and, in some cases, mycelia embedded needles. The bacterial development observed in one of the studied needles was interpreted as a contamination by an enchytraeid, the tegument of which probably carried propagules and deposited them at the inside of the needle. A relationship between bacterial development and fungal death at the inside of the needle was hypothesized but not ascertained.

The observed fungal successions were in full agreement with the observations by Kendrick & Burges (1962), to the exception of the time lapse they indicated for the fructification of *Lophodermium pinastri* and *Verticicladium pinastri*. In the study site, located much more southerly compared to England, these two fungi fructified within at most two months, i.e. much more rapidly than the 6-8 months these authors indicated. However, Soma & Saito (1979), in a study on the decomposition of needles of *Pinus thunbergii* in climate conditions comparable to the present study, found 6-9 months for the maturation of ascocarps of *Lophodermium pinastri*. For the sake of comparison, it should be also noted that *Ceuthospora pinastri* was erroneously called *Fusicoccum bacillare* by Kendrick & Burges (1962), as this could be ascertained from their figures, and that my L₁ layer corresponded to their L layer, while my L₂ layer corresponded to their F₁ layer.

## Decaying twigs

A 2-yr-old twig was sectioned, its age being estimated from the succession of early- and latewood in the observed transversal sections.

Black pycnidia were present around the twig, protruding through peeled off bark and epidermis (Fig. 30). At least in some zones which were kept untouched, parenchymatous tissues were attacked by networks of hyaline and melanised hyphae, into which several species were seen but could not be identified. This was the case for the medulla (Fig. 31), the cortex (Fig. 32), medullary rays (Fig. 33) and vascular rays (Fig. 34). Figure 35 shows the passage from the intact protoxylem (compare with Figure 36) to the intact metaxylem, passing through a parenchymatous zone (medullary ray) which was the seat of an intense development of melanised hyphae.

When cells were attacked, cellulose became brown and lignin remained transparent. At this early stage of branchwood decay, the penetration of tracheids by hyphae from vascular rays was incipient (Fig. 37). Parenchymatous cells of vascular rays communicate freely with metaxylem tracheids through enlarged pits without torus (Fig. 34). Thereby the fungal colonisation of branchwood was both centripetal (through the cortex) and centrifugal (through the medulla) and was mainly mediated by vascular rays. At this stage, no internal development of ligninolytic basidiomycetes was registered.
<figure><img src="image_7.png" /><figcaption>Fig. 34: Two-yr-old pine twig, longitudinal section showing a vascular ray (BMSL); Fig. 35: Idem, longitudinal section showing, from top to bottom, metaxylem, medullar parenchyma, protoxylem (BMSL); Fig. 36: Idem, longitudinal section showing a reticulated tracheid of the protoxylem (CP); Fig. 37: Idem, longitudinal section showing the passage of melanised hyphae from a vascular ray to adjoining tracheids (BMSL); Fig. 38: Piece of bark, section showing micro-algae in tetrads (BMSL); Fig. 39: Idem, melanised hyphae at the surface of the epidemis (CP); Fig. 40: Idem, brown deposits on walls of bark cells (BMSL); Fig. 41: Idem, detail (BMSL)</figcaption></figure>

CP = phase contrast; BMSL = methyl blue-Sudan III-lactophenol coloration; r.v. = vascular rays; h.m. = melanised hyphae; c.a. = micro-algal colonies; length of segment = 50 µm
## Bark fragments

These bark fragments were detached from the tree. Their surface was covered with unidentified pleurococcoid micro-algae, cells of which were ~2 µm (Fig. 38). When epidermis was still present, it was covered with a network of melanised hyphae (Fig. 39). The walls of dead cells of bark (suber) were lightly brown and covered with dark brown deposits (Figs. 40 & 41). The lumen of some bark cells was near entirely filled with light brown masses (Fig. 42). No fungal development was visible.

## Living and decaying moss (*Pseusceropodium purum*)

Intact leafy shoots of *Pseudoscleropodium purum* were the dominant component of the L₁ layer, showing typical anatomical features of living leaves (Fig. 43). Like other Bryales *Pseusceropodium purum* foliar limbs are made of a single layer of photosynthetic cells, without any lamella (de Ferré, 1963). I observed on some moss leaves that the photosynthetic tissue was directly attacked by dematiaceous fungi crawling at the surface of foliar limbs (Fig. 44).

## Fauna

### Inventory

Table 3 shows that oribatid mites were the most important invertebrate group, both in density (46 specimens) and in biovolume (0.24 µL < V < 0.98 µL). It should be noticed that a unique specimen of the springtail *Pogonognathellus flavescens* (Tullberg) occupied a biovolume comprised between 0.07 and 0.28 µL. This is exemplary of huge variations which could occur in soil animal biovolume (or biomass) when a big animal appears or disappears from a prospected area.

Concerning nematodes, as well as for the rest of microfauna, it must be noticed that my method does not allow collecting adequately tiny transparent animals such as nematodes, rotifers and protozoa. Only one nematode (of a big size for this group) was observed and studied, which is not apportioned to their actual abundance. However, I can assure with a reasonable degree of confidence that they were not numerous in the L₁ layer, given that, with the same method, many more specimens were found in underlying layers, as well as many cyst amoeba.

### Enchytraeid worms

Although they could not be identified at the species level, given the methods used for fixing and preserving the studied material, all specimens present in the studied samples probably pertained to the genus *Cognettia* (A. Albrecht, personal communication).
<figure><img src="image_8.png" /><figcaption>Fig. 42: Piece of bark, section showing light brown spherical masses filling the lumen of some bark cells (BMSL); Fig. 43: Leaving leaf of the moss *Pseudoscleropodium purum* (CP); Fig. 44: Melanised hyphae crawling at the surface and invading moss leaf cells; Fig. 45: Items carried on the tegument of enchytraeids, micro-algae in tetrads and silt-size quartz grains (CP); Fig. 46: Idem, melanised fungal hyphae with smooth walls (CP); Fig. 47: Idem, pollen grain of pine, intact (CP); Fig. 48: Idem, moss leaf fragment of *Pseudoscleropodium purum* (CP); Fig. 49: Idem, springtail faeces (CP)</figcaption></figure>

CP = phase contrast; BMSL = methyl blue-Sudan III-lactophenol coloration; h.m. = melanised hyphae; g.q. = quartz grains; a. = micro-algae; d. = faeces; G.P. = pollen grains; length of segment = 50 µm
<table><caption>**Table 3.** Taxonomic name, abundance and estimated biovolume of animals found in the L₁ layer</caption><thead><tr><th>Taxonomic groups and species</th><th>Abundance</th><th>Upper estimate of biovolume (µL)</th><th>Lower estimate of biovolume (µL)</th></tr></thead><tbody><tr><td>Nematodes</td><td>1</td><td>0.69 10<sup>-3</sup></td><td>0.17 10<sup>-3</sup></td></tr><tr><td><i>Prionchulus punctatus</i></td><td>1</td><td>0.69 10<sup>-3</sup></td><td>0.17 10<sup>-3</sup></td></tr><tr><td>Enchytraeids</td><td>7</td><td>0.22</td><td>0.06</td></tr><tr><td>Oribatid mites</td><td>46</td><td>0.98</td><td>0.24</td></tr><tr><td><i>Platynothrus peltifer</i></td><td>22</td><td>0.49</td><td>0.12</td></tr><tr><td>Chamobates sp.</td><td>3</td><td>0.61 10<sup>-1</sup></td><td>0.15 10<sup>-1</sup></td></tr><tr><td>Parachipteria punctata</td><td>3</td><td>0.55 10<sup>-1</sup></td><td>0.14 10<sup>-1</sup></td></tr><tr><td>Oppia sp.</td><td>3</td><td>0.13 10<sup>-1</sup></td><td>0.03 10<sup>-1</sup></td></tr><tr><td>Oribatula tibialis</td><td>3</td><td>0.26 10<sup>-1</sup></td><td>0.06 10<sup>-1</sup></td></tr><tr><td>Camisia spinifer</td><td>2</td><td>0.97 10<sup>-1</sup></td><td>0.24 10<sup>-1</sup></td></tr><tr><td>Ceratoppia bipilis</td><td>2</td><td>0.12</td><td>0.03</td></tr><tr><td>Oppiella tricarinata</td><td>2</td><td>0.60 10<sup>-2</sup></td><td>0.15 10<sup>-2</sup></td></tr><tr><td>Oppiella subpectinata</td><td>2</td><td>0.17 10<sup>-1</sup></td><td>0.04 10<sup>-1</sup></td></tr><tr><td>Phthiracarus sp.</td><td>1</td><td>0.42 10<sup>-1</sup></td><td>0.10 10<sup>-1</sup></td></tr><tr><td>Carabodes femoralis</td><td>1</td><td>0.19 10<sup>-1</sup></td><td>0.05 10<sup>-1</sup></td></tr><tr><td>Adoristes ovatus</td><td>1</td><td>0.26 10<sup>-2</sup></td><td>0.10 10<sup>-2</sup></td></tr><tr><td>Minhuntozetes semirufus</td><td>1</td><td>0.40 10<sup>-2</sup></td><td>0.10 10<sup>-2</sup></td></tr><tr><td>Miscellaneous mites</td><td>4</td><td>0.49 10<sup>-1</sup></td><td>0.12 10<sup>-1</sup></td></tr><tr><td>Uropodidae</td><td>3</td><td>0.44 10<sup>-1</sup></td><td>0.11 10<sup>-1</sup></td></tr><tr><td>Bdellidae</td><td>1</td><td>0.45 10<sup>-2</sup></td><td>0.11 10<sup>-2</sup></td></tr><tr><td>Spiders</td><td>2</td><td>0.49 10<sup>-1</sup></td><td>0.08 10<sup>-1</sup></td></tr><tr><td>Springtails</td><td>9</td><td>0.30</td><td>0.07</td></tr><tr><td><i>Parisotoma notabilis</i></td><td>8</td><td>0.16 10<sup>-1</sup></td><td>0.04 10<sup>-1</sup></td></tr><tr><td><i>Pogonognathellus flavescens</i></td><td>1</td><td>0.28</td><td>0.07</td></tr><tr><td>Fly larvae</td><td>7</td><td>0.38</td><td>0.10</td></tr><tr><td>Cecidomyidae</td><td>5</td><td>0.25</td><td>0.06</td></tr><tr><td>Sciaridae</td><td>1</td><td>0.11</td><td>0.03</td></tr><tr><td>Chironomidae</td><td>1</td><td>0.17 10<sup>-1</sup></td><td>0.04 10<sup>-1</sup></td></tr><tr><td>Corpses</td><td>4</td><td></td><td></td></tr><tr><td>Unidentified arthropod</td><td>1</td><td></td><td></td></tr><tr><td>Unidentified oribatid mite</td><td>1</td><td></td><td></td></tr><tr><td><i>Platynothrus peltifer</i></td><td>1</td><td></td><td></td></tr><tr><td><i>Phthiracarus sp.</i></td><td>1</td><td></td><td></td></tr></tbody></table>

* Transport on the tegument

The direct examination of enchytraeids showed that their tegument was covered with sticky mucus onto which a variety of items adhered and could identified under the light microscope: pleurococcoid green micro-algae (Fig. 45), melanised fungal hyphae with smooth (Fig. 46) or echinulate cell walls, and fungal spores. Fungal hyphae seemed to be intact and of a length indicating that they were more probably viable and thus prone to disperse the fungus. Pollen grains were also found (Fig. 47), as well as leaf fragments of *Pseudoscleropodium purum* (Fig. 48), springtail faeces (Fig. 49), nematodes (Fig. 50) and numerous quartz grains of silt size (Fig. 45). The presence of silt particles on the tegument of enchytraeids was the only trace of mineral uplift I saw in the L₁ layer. These observations testified for the important carrying activity exerted by enchytraeids: they disseminate micro-algae, fungi and bacteria and effect mineral uplift. Given the thickness of organic layers (8 cm) the presence of quartz grains of silt size in the L₁ layer indicated the important vertical movements effected by these animals.
<figure><img src="image_9.png" /></figure>

Fig. 50: Items carried on the tegument of enchytraeids, nematode (CP); Fig. 51: Gut contents of enchytraeids, observed by transparency, pollen grain of pine, collapsed (CP); Fig. 52: Idem, moss leaf fragment of *Pseudoscleropodium purum* (CP); Fig. 53: Idem, long fragments of fungal melanised hyphae (CP); Fig. 54: Idem, intestinal microflora (CP); Fig. 55: Idem, cyst amoeba (*Nebela collaris*) (CP); Fig. 56: Idem, faecal material of oribatid mites, with fungi (degraded) and bacteria (CP); Fig. 57: Idem, faecal material of oribatid mites, with melanised, highly compacted fungal hyphae (CP)

CP = phase contrast; nem. = nematode; G.P. = pollen grains; h.m. = melanised hyphae; c.b. = bacterial colonies; length of segment = 50 µm
• Feeding activity

Gut contents of enchytraeids were observed by transparency: pollen grains (Fig. 51), fragments of leaves of *Pseudoscleropodium purum* (Fig. 52), non-fragmented melanised hyphae (Fig. 53). Bacterial colonies, which seemed living according to their neat and refringent contour under phase contrast, were attributed to intestinal microflora (Fig. 54). Cyst amoebae were also sometimes found, such as *Nebela collaris* (Ehrenberg) (Fig. 55). Mite faeces were also ingested. In this case fungal material seemed much more degraded, mixed with bacteria (Fig. 56) or highly compacted (Fig. 57). The necessity of a double digestion for the degradation of melanised fungal walls must be highlighted, which is reminiscent of observations made by Reisinger (1972) on the mite *Tyrophagus putrescentiae* (Schrank).

### Nematodes

A single specimen, belonging to *Prionchulus punctatus* (Cobb) (Mononchidae), was found, but we expect nematodes were much more abundant. Their small size and transparency allowed them, together with other microfauna, to escape my observations with a dissecting microscope of the material fixed into ethyl alcohol. The observed specimen was considered as a predator (Arpin, 1980). By transparency, the degradation of the ingested material could be followed. In the anterior part of the intestine, I found mastax of rotifers (Fig. 58), thereby indicating that these animals served as food, only hard chitinous parts remaining undigested. Bacterial cells had slightly fuzzy contours, were poorly refringent under phase contrast, but they were still individualised. The medium part of the intestine showed more degraded bacteria, with a fuzzier contour, together with debris of melanised fungal spores (Fig. 59). In the posterior part, just before anus, numerous bacteria, highly compacted and with a hardly discernible contour, could be observed (Fig. 60). It must be noticed that from the examination of its gut contents the food diet of this 'predatory' nematode was mainly made of bacteria, which were progressively degraded during intestinal transit. Similar observations, using transmission electron microscopy, were done by Arpin & Kilbertus (1981) on the nearby species *Prionchulus muscorum*.

### Oribatid mites

The species dominant in number and biomass (as ascertained by biovolume, see Table 3) was *Platynothrus peltifer* (Koch), which was represented by tritonymphs and adults. When isolated by dissection, the intestine showed by transparency several food boluses, similar in shape (oval) and size (50-100 µm) to oribatid faecal pellets, and bacterial colonies in special pouches (caecums) freely communicating with the intestinal lumen (Fig. 61). One or several colonies of intestinal microflora were found in all specimens, whether juvenile or adult, starving or feeding, which were dissected. All bacterial colonies living in the intestine of *Platynothrus peltifer* seemed identical morphologically. The ingested material was made of fungal hyphae, cut into pieces (contrary to what was observed in
<figure><img src="image_10.png" /></figure>

Fig. 58: Gut contents of the nematode *Prionchulus punctatus* (Mononchidae) observed by transparency, anterior intestine showing rotifer mastax, bacteria, unidentifiable organic matter (walls of melanised hyphae?) (CP); Fig. 59: Idem, medium intestine showing degraded bacteria and broken fungal spores (CP); Fig. 60 (x 400) Idem, posterior intestine showing strongly degraded bacteria (CP); Fig. 61: Gut contents of the oribatid mite *Platynothrus peltifer* observed after dissection, intestinal microflora (CP); Fig. 62: Idem, fungal spores and melanised hyphae (CP); Fig. 63: Gut contents of the oribatid mite *Adoristes ovatus* observed by transparency, showing three stages in the formation of faecal pellets (CP); Fig. 64: Gut contents of the springtail *Parisotoma notabilis*, degraded melanised hyphae and bacteria (CP); Fig. 65: Gut contents of fly larvae (Sciaridae) observed by transparency, leaves of the moss *Pesudoscleropodium purum* (CP)

CP = phase contrast; m.r. = mastax of rotifers; c.b. = bacterial colonies; h.m. = melanised hyphae; sp. = fungal spores; length of segment = 50 µm
enchytraeids), mostly melanised (dematiaceous), and spores (Fig. 62). Fungal cell walls seemed not
too degraded.

A similar food diet was found in *Ceratoppia bipilis* (Herman), *Camisia spinifer* and *Adoristes ovatus*. A photograph was taken by transparency through a specimen of the latter species, showing three stages of compaction of fungal hyphae which occurred during the formation of faeces (Fig. 63).

Springtails

Gut contents of *Parisotoma notabilis* (Schäffer) were, here too, formed of fungal material (Fig. 64) but, contrary to *Platynothrus peltifer*, bacteria were present in the food bolus and fungal hyphae were degraded, suggesting that a decaying and not a living fungal colony was ingested, probably coming from underlying layers (faecal material?).

Fly larvae

Only results about *Sciaridae* and *Chironomidae* will be presented, since not any solid particles were
found in intestines of *Cecidomyidae*.

Sciarid larvae ingested mainly fragments of the moss *Pseudoscleropodium purum* (Figs. 65 & 66) but I found also melanised fungal hyphae, which had been ingested without being cut into pieces (Fig. 67), and pollen grains of *Pinus sylvestris* (Fig. 68). It must be noticed that fungal hyphae seemed to be ingested in the same manner as enchytraeids, without any previous fragmentation.

Fungal melanised hyphae were found in the intestines of chironomid larvae, together with hyaline hyphae. In the latter case, hyphae appeared compacted, with fuzzy contours, in way to be degraded (Fig. 69). Brown masses of organic matter (humus), of unknown origin, were also found (Fig. 70). It must be highlighted that the degradation of hyaline and melanised fungal hyphae was hardly comparable, melanin making hyphal walls of dematiaceous fungi resistant to gut enzymes (Reisinger, 1972).

Animal faeces

Only faeces of oribatid mites were observed in the L₁ layer. They were few abundant and deposited at the surface of needles and bark pieces. They appeared green in freshly collected material. Examination of faecal material under the light microscope showed that oribatid faeces contained decaying fungal material with numerous micro-algae in vegetative growth, as ascertained by preferential staining of chromosomes by methyl blue (Fig. 71).

Similar micro-algae were found in gut contents of *Platynothrus peltifer*, although in few
<figure><img src="image_11.png" /></figure>

Fig. 66: Gut contents of fly larvae (Sciaridae), observed by transparency, leaf fragment of the moss *Pseudoscleropodium purum* (CP); Fig. 67: Idem, long fragments of melanised fungal hyphae (CP); Fig. 68: Idem, pollen grain of pine (CP); Fig. 69: Gut contents of fly larvae (Chironomidae) observed by transparency, hyaline hyphae, compacted and in way to be degraded (CP); Fig. 70: Idem, unidentifiable brown organic matter (melanised fungal walls?) (CP); Fig. 71: Micro-algal development (metaphases) in oribatid faeces (BMCL); Fig. 72: Unidentified cysts in an arthropod corpse (CP); Fig. 73: Development of melanised fungal hyphae in a corpse of *Platynothrus peltifer* (observed by transparency) (CP)

CP = phase contrast; BMCL = methyl blue-chloral-lactophenol; h.m. = melanised fungal hyphae; G.P. = pollen grains; r.d.h.h. = dense network of hyaline hyphae; al. = micro-algae; met. = metaphases; ky. = cysts; length of segment = 50 µm
abundance. It is possible that micro-algal development started by some algal cells which were ingested
then excreted in living stage in the faeces, thereafter developing in this favourable environment.
Contrary to animal guts, into which fungal walls of melanised hyphae were still intact, the same
material was in way to be degraded in faeces invaded by micro-algae.

Animal corpses

All corpses found in the L₁ layer were arthropod cadavers. A corpse of *Platynothrus peltifer* was shown to be invaded by a dense network of melanised hyphae (Fig. 72), while a corpse of an unidentified arthropod was filled with cysts of unknown origin (Fig. 73).

Conclusions about the L₁ layer

The decomposition of pine needles in the L₁ layer was mainly ensured by fungi. Even though the external development of fungi was but feeble, all needles were invaded by fungal hyphae which penetrated internal tissues (mesophyll, transfusion tissue) and replaced progressively the cytoplasm of plant cells. When needles were colonised on the tree (by *Lophodermium pinastri*, *Lophodermella* or *Ceuthospora pinastri*) part of this process had already started before needles fell on the ground.

Needles in the most advanced stage of decomposition (probably in contact with the underlying
level L₂) were characterised by the internal development of *Verticicladium trifidum*. Blackening of
hypodermic call walls under the influence of this fungus is worth to notice. Needle collapse was
associated with the disappearance of cell cytoplasm, in particular in the mesophyll.

Similarly, parenchymatous tissues of pine twigs were colonised by fungi. The attack of wood
was only incipient, some hyphae penetrating tracheids through vascular rays.

The impact of fauna seemed limited to grazing of fungi which grew externally on the decaying
plant material. However, it must be noticed that leaves of the moss *Pseudoscleropodium purum* were a
major food resource for sciarid larvae and to a lesser extent for enchytraeids. The transport of bacteria,
fungal propagules (hyphae, spores) and mineral grains by the latter animal group was also noticeable
in this superficial layer, exemplifying the importance of invertebrates in the early microbial
colonisation of pine litter. Bacterial development at the inside of a pine needle tunnelled by
enchytraeids was also indicative of the role of soil animals as carriers of bacteria.
THE L₂ LAYER: DECAYING NEEDLES AND DEAD MOSS

<figure><img src="image_12.png" /><figcaption>**Figure 74.** The L₂ layer before fixation and transport to the laboratory</figcaption></figure>

Table 4 shows the respective volume of the gross components of the L₂ layer. Compared to the overlying L₁ layer, the L₂ layer was characterised by a greater diversity of plant material and a measurable amount of animal faeces.

<table><caption>**Table 4.** Volume of the different components of the L₂ layer sorted under the binocular microscope</caption><thead><tr><td>Litter components</td><td>Volume (µL)</td></tr></thead><tbody><tr><td>Pine wood</td><td>5100 ± 100</td></tr><tr><td>Moss</td><td>2600 ± 100</td></tr><tr><td>Pine bark</td><td>1700 ± 100</td></tr><tr><td>Pine needles</td><td>1700 ± 100</td></tr><tr><td>Animal faeces</td><td>220 ± 10</td></tr><tr><td>Pine twigs</td><td>200 ± 10</td></tr><tr><td>Bracken fronds</td><td>80 ± 10</td></tr><tr><td>Pine mycorrhizae</td><td>70 ± 10</td></tr><tr><td>Male flowers of pine</td><td>22 ± 2</td></tr><tr><td>Pine euphylls</td><td>12 ± 2</td></tr><tr><td>Living fauna</td><td>0.8 – 3.4</td></tr><tr><td>Total</td><td>11700 ± 100</td></tr></tbody></table>
Litter fall corresponding to the L₂ layer could be dated from previous December-January to previous June. The earlier date was deduced from the presence of bracken litter, *Pteridium aquilinum* (L.), in the form of leaflets and secondary and tertiary rachises detached from the fronds which currently fall in winter. The later date was deduced from the presence of pine euphylls, the fall of which currently occurs in June. Thus the L₂ layer embraced a period of time four times longer than the overlying L₁ layer.

The amount of moss material (22% of the total volume) was much less than in the L₁ layer (77%). However, this estimate must be amended because of the presence of a 5-yr-old branch of pine, the volume of which was estimated to 44% of the total volume of the sample. After discarding branchwood, the volume of dead moss was 39%, which still indicated a pronounced decrease compared to L₁. This should not be interpreted as resulting from a high decay rate of moss compared to pine needles, but rather to a low primary production of moss in winter and spring. At this stage, leafy stems of *Pseudoscleropodium purum* were deprived of pigment and invaded by an abundant mycelium but seemed still intact, at least to the naked eye (Fig. 74).

As in the previous layer, dating of the moss fraction was difficult. Kilbertus (1968a, b) showed that *Pseudoscleropodium purum* remained green for a year and a half then senesced and became brown for an additional three years, before decaying. At first sight this contradicts my observations on pine and bracken litter in the studied sample. Such a marked disagreement could be explained by the fact that (i) pine needles, when falling on the ground, penetrated to some extent through the moss carpet, (ii) moss continued to grow upwards after needles had fallen, and tended to cover them up during spring and summer months i.e. just previous to sampling which took place in August. However, this does not explain why dead foliage of *Pteridium aquilinum* (not likely to penetrate through moss) was found in the same layer than dead parts of *Pseudoscleropodium purum*. The present study cannot answer this question, but it should be noticed that similar discrepancies were found between my field observations and experiments on decomposition rates of pine needles performed by Kendrick & Burges (1962), as mentioned above. Guittet (1967) showed that the 'duration' of the L₂ layer was thrice that of the L₁ layer in Scots pine stand, which compares well with my observations. Unfortunately this author could not adequately date its litter samples.

Data on animal faeces presented in Table 4 did not embrace the whole faecal material. Although in much higher abundance in L₂ than in the overlying L₁ layer, faecal pellets of enchytraeids and microarthropods were too isolated and could not be taken with forceps, and thus their biovolume could not be measured adequately with my method. Rather, I estimated the volume of faeces of macroinvertebrates such as earthworms, woodlice and millipedes.

Voids were important (77%), although slightly less than in the overlying layer (80%), showing the absence of packing of pine and bracken litter through the moss carpet. The biovolume of fauna, mostly mesofauna, was feeble (0.8-3.4 µL) although slightly higher than in L₁ (0.4-2 µL).

Externally L₂ pine needles differed to a great extent from those found in L₁. They were black-coloured, often covered, as the whole material present in L₂, by a loose weft of mycelia, and most of them were collapsed (Fig. 74). Traces of previous colonisation by *Lophodermium pinastri* and *Ceuthospora pinastri* were still visible, but *Verticicladium trifidum* was dominant: many needles were covered by a net of its conidiophores. The folds of collapsed needles, where conidiophores of *Verticicladium trifidum* formed a dense network, were the main niche of nematodes, rotifers, ciliates and cyst amoebae, which could not be adequately collected extensively because of their small size and
transparency (a limit of my method, which was not convenient for microfauna). Brown rhizomorphs of *Marasmius androsaceus* Fries were seen protruding from some needles, which were in way to be discoloured. Similar observations of needle discoloration by *Marasmius androsaceus* were made by Soma & Saitô (1979) on *Pinus thunbergii* Parl., by Gourbière (1981) on *Abies alba* Mill., and by Latter (1977) on *Calluna vulgaris* Salisb. and several heathland species. This basidiomycete is a white rot (Hintikka, 1970). Its rhizomorphs have been considered by Gourbière (1981) to penetrate then to colonize coniferous needles through internal mycelia. In the present sample, rhizomorphs were short (a few cm), their width decreased then they ramified at distance from colonised needles, without linking them. They were interpreted as issuing from needles and not entering them. Mitchell & Millar (1978) showed that the cuticle of pine needles was penetrated by thin hyphae ramifying from rhizomorphs appressed on the cuticle and that rhizomorphs protruded from the needles by lifting out hypodermis, epidermis and cuticle. Although I did not observe any fungal rhizomorph penetrating the needles, I cannot discard that direct penetration by rhizomorphs of *Marasmius androsaceus* could have occurred previously. The fact that this fungus was absent from L₁ suggests that infection took place during previous winter or spring but was not extended to summer.

Some needles exhibited fungal sclerotia of particular toughness and shape, which could not be identified by lack of fructifications.

The action of fauna was prominent in the L₂ layer. In addition to macroinvertebrate activity, traced from the deposition of faeces (2% of the total volume of solid matter), most animal activity taking place at the time of sampling was attributed to enchytraeids. All pine needles showed signs of biting or penetration by these animals, and a needle had even been fixed with an animal penetrating it. Some other needles showed breaks, which could not be attributed to enchytraeids but rather to macroinvertebrates which deposited their faeces at this level. Most enchytraeids were retrieved in decaying leafy shoots of the moss *Pseudoscleropodium purum*, at the inside of fungal sheaths embedding stems and leaves. I noticed the behavioural tendency of these worms to penetrate any kind of decaying plant material. Several specimens were retrieved between bark and wood, among their faecal deposits, in the decomposing pine branch which was studied in the L₂ sample. Other deposits of enchytraeid faeces were observed at the surface of bark pieces, between needles when still assembled by pairs and appressed to each other, between stems and leaves of moss, or in fluffy masses invaded by mycelial wefts. Faeces of mites and springtails were more abundant than in L₁. They were of a dark brown colour and most of them were found isolated at the surface of needles.

Mycorrhizal fine roots of pine were not very abundant at this superficial level (0.6% of total volume of solid matter), being preferentially observed in F₂ and H₁ (Table 1). They were embedded in white-grey mycelial wefts of woolly aspect, quite identical to those observed around well-decayed pine needles, leafy shoots of mosses and enchytraeid faecal deposits.

The abundance of bark was also noticeable, with a biovolume comparable to that of pine needles (14.5%), as was already true in L₁. I want to notice that bark, a prominent component of pine litter, has been unexpectedly neglected in decomposition studies, if we except Kuhlman (1969, 1970).

Decaying pine needles
As for the overlying L₁ layer, only a small number of representative types will be described, although a wide variety of needles were sectioned (15 different morphological types).

*Needles attacked by Verticicladium trifidum*

Compared to L₁, the attack was much more advanced. Needles turned to full black over their whole surface. Sectioned needles showed that the contact zone between hypodermis and mesophyll was the main seat of blackening (Fig. 75): a thin black opaque layer, not made of fungal hyphae, was seen blanketing the hypodermis. Blackening diffused through cell files of the mesophyll, still recognizable at this stage of decay in the absence of living hyphae.

Cell walls of internal tissues were destroyed to a higher extent than in L₁. Endodermis was degraded, as well as the external part of transfusion tissue (Fig. 76). The degradation of lignin was effective, as ascertained by the presence of more or less identifiable bordered pits (areolae) within a mass of tracheid wall remains. The attack was more recent (less pronounced) at the level of vascular tissues: lignified walls were still intact, but I observed some living hyphae penetrating tracheids of the metaxylem through bordered pits, probably after disappearance of the torus (Fig. 77). Long hyaline hyphae of *Verticicladium trifidum*, stained blue by methyl-lactophenol, were observed running along vessels of secondary wood.

Although *Verticicladium trifidum* was often reported as a primary coloniser of fallen pine needles, mycological studies did not register its paramount importance in the decay process. Practically nothing was said of hyaline hyphae invading first living tissues, the cytoplasm of which was destroyed, then other tissues where cellulose and lignin were destroyed in turn. According to my observations, *Verticicladium trifidum* was always associated with the presence of black stromata, which seemed typical of this fungus. Mitchell & Millar (1978) observed that the bleaching (?) of pine needles was associated with the development of black stromata, to which they attributed the degradation of cellulose. Conversely, and more in conformity with my own observations, Lehmann & Hudson (1977) attributed to *Verticicladium trifidum* both blackening and softening of pine needles, the latter process being probably due to the degradation of lignin. Kendrick & Burges (1962) considered this fungus to be responsible for needle blackening and they demonstrated on a quantitative basis that sporulation (see conidiophore on Fig. 6) occurred in summer. They showed that it produced two kinds of hyphae, hyaline hyphae penetrating internal tissues, and melanised hyphae penetrating epidermis and hypodermis and thought to be responsible for needle blackening. The present study showed that at the inside of pine needles the mycelium of *Verticicladium trifidum* was only made of hyaline hyphae, melanised hyphae being entirely transformed in melanised stromata. Only hyaline hyphae were seen invading host cells and digesting their cytoplasm in the vicinity of stromata, such as for instance in stomatal guard cells (Figs. 21 & 22). Melanised stromatic nodules were seen from place to place in internal tissues, stemming from hyaline hyphae (Fig. 78). Such results conform to observations made in culture (Hughes, 1951; Gremmen, 1960) or by directly observing decaying needles (Hughes, 1951).

*Needles attacked by Marasmius androsaceus*
<figure><img src="image_13.png" /><figcaption>Fig. 75: Needle attacked by *Verticicladium trifidum*, longitudinal section showing epidermis, hypodermis, mesophyll, endodermis, transfusion tissue and perivascular fibres (BMSL); Fig. 76: Idem, detail of transfusion tissue showing partial degradation of cell walls (CP); Fig. 77: Idem, detail of metaxylem showing penetration of tracheids through bordered pits (arrows) (BMSL); Fig. 78: Idem, detail showing melanised stromata within transfusion tissue (CP); Fig. 79: Needle attacked by *Marasmius androsaceus*, brown rhizomorph (CP); Fig. 80: Idem, section through a zone from which started a rhizomorph, showing mesophyll, perivascular fibres, protoxylem and metaxylem (CP); Fig. 81: Idem, section through a fold of a collapsed needle showing composite bacterial colonies (CP); Fig. 82: Idem, showing a quartz grain of silt size covered with minute bacteria (BMSL)</figcaption></figure>

CP = phase contrast; BMSL = methyl blue-Sudan III-lactophenol coloration; c.n. = black opaque layer; t.m. = cell files of the mesophyll; ar. = areolae; ps. = pseudoparenchyma; b. = bacteria; length of segment = 50 µm
This basidiomycete formed brown, leathery, flat rhizomorphs (Fig. 79), the anatomy of which was conform to descriptions given by Macdonald (1949) and Townsend (1954). According to criteria of maturity, such as the development of the internal lumen (responsible for flatness), rhizomorphs found in the L₂ layer were ageing. This is an additional argument for dating fungus infestation by *Marasmius androsaceus* to last autumn/winter.

In the vicinity of rhizomorph points of output the mesophyll appeared totally degraded, being replaced by fungal pseudoparenchyma (Fig. 80). Lignified walls of tracheids remained intact, but bacteria could be observed at the inside of tracheids with bordered pits of the metaxylem. This is another indication of ancientness of the colonisation process.

Needles colonised by *Marasmius androsaceus* were collapsed, due to disappearance of the mesophyll. Numerous folds were observed, with bacterial and micro-algal colonies, sometimes formed by several species in tight assembly (Fig. 81). Quartz grains, probably transported by enchytraeids or by earthworms, were also found in these microcosms: they were covered with minute bacteria lodging in irregularities (hollows) of the crystal surface (Fig. 82).

*Marasmius androsaceus* was suspected for a long time to be a lignin degrader (Lindeberg, 1944), which was confirmed by histological studies on infested needles (Mitchell & Millar, 1978; Mitchell et al., 1978a), but these authors did not show, as in the present study, any penetration of tracheids by this fungus. Bacteria, which were scarcely represented within tracheids, probably came from further contamination and did not show any sign of degradation of tracheid walls. However, the impact of the fungus on cellulose was prominent in the zone of rhizomorph rooting. I cannot discard the possibility of an erroneous identification of the fungus, which was done only through rhizomorphs and not through carpophores (which were absent from my sample).

*Needles attacked by unidentified fungi*

Some needles showed one or more black sclerotia at their surface. These needles were at a more advanced stage of decomposition (they were internally colonised by bacteria) and the action of the fungus previous to bacterial colonisation could not be ascertained.

Another species of fungus formed white excrescences at the surface of some needles. This fungus developed at the inside of needles, in mesophyll and transfusion parenchymatous tissues in which it formed long strands made of fibrous hyphae with thick wall and narrow lumen (Fig. 83). Hyphae of this fungus did not penetrate host cells, but rather its fibrous strands pulled aside host tissues, cells of which were squashed but kept intact their walls (Fig. 84). Externally, the fungus developed a network of fibrous hyphae forming the so-called white excrescences (Fig. 85). At this stage, needles were often embedded in a thick sheath made of dead melanised hyphae, which was colonised by bacteria (Fig. 86) and micro-algae (Fig. 87). This fungal-bacterial-micro-algal sheath was lifted from place to place by the development of white excrescences, which were able to pierce it and develop externally in the form of white, tough, smooth white masses. The external network of melanised hyphae was reminiscent of the 'network stage' which has been described by many authors (Kendrick & Burges, 1962; Hayes, 1965; Lehmann & Hudson, 1977; Mitchell et al., 1978a; Mitchell & Millar, 1978; Soma & Saitô, 1979; Gourbière, 1981). It was relict of an earlier stage of fungal
<figure><img src="image_14.png" /><figcaption>Fig. 83: Needle attacked by an unidentified fungus with white excrescences, longitudinal section showing transfusion tissue (BMSL); Fig. 84: Idem, mesophyll (BMSL); Fig. 85: Idem, dense fibrous mass at the surface of a needle (BMSL); Fig. 86: Idem, network of dead melanised hyphae with bacteria (BMSL); Fig. 87: Idem, with micro-algae (BMSL); Fig. 88: Needle with a development of bacteria, longitudinal section showing protoxylem (BMSL); Fig. 89: Idem, mesophyll (BMSL); Fig. 90: Needle with a development of bacteria and micro-algae, tangential section through epidermis (BMSL)</figcaption></figure>

BMSL = methyl blue-Sudan III-lactophenol coloration; h.h. = hyaline hyphae; h.m. = melanised hyphae; length of segment = 50 µm
colonisation, probably under moister atmosphere during previous winter or spring. The dry
atmosphere of summer months might have limited the external development of fungi in L₁ (the past of
L₂) while the internal development of *Verticicladium trifidum* did not seem to be affected by dry air,
higher temperature allowing it to grow and fructify rapidly.

*Needles with bacterial development*

These needles were always broken or nibbled, indicating a previous attack by fauna. Bacteria seemed
to attack preferentially fungal hyphae. In needle tissues where fungal attack by *Verticicladium trifidum*
was still incipient and did not degrade host cells, such as xylem, hyphae of this fungus could be
morphologically identified, with bacteria at their inside (Fig. 88). In the mesophyll, in which previous
attack by fungi was more pronounced, melanised fungal walls were degraded. They were probably
relicts of another (unidentified) dematiaceous fungus. Melanised walls appeared fragmented and
agglomerated in coarse masses (Fig. 89). Studying at the right scale this process of transformation
(humification?) of melanised fungal cell walls and establishing the role of bacteria would need the use
of electron microscopy, which was out of the scope of my study.

*Needles with bacterial and micro-algal development*

These needles have been also previously broken by soil invertebrates. I interpreted this stage as a more advanced stage of decomposition compared to bacteria only. Needles were markedly collapsed, with numerous folds. Epidermis and hypodermis were replaced by a melanised fungal stroma, dead at this stage (Fig. 90), which was covered by a dense network of dead melanised hyphae, morphologically identical to the abovementioned 'network stage', with numerous bacteria.

The inside of needles was in an advanced stage of decomposition, under the influence of an
intense bacterial development. Areolae from bordered pits were often the only trace of pre-existing
tracheids (Fig. 91). The development of micro-algal colonies was visible at the inside of needles. I
observed Chlorophycae (Fig. 92), chloroplasts of which were coloured in violet by methyl blue-Sudan
III (same colouration as in green needles and moss leaves), and Cyanophyceae of *Chroococcus* type
(Fig. 93), the cytoplasm of which was uniformly coloured in blue without any nuclear or plastidial
inclusion.

Some needles at this stage of decomposition showed a development of protozoa, as
ascertained by the presence of numerous cysts (Fig. 94). Protozoan trophozoits were not observed and
I was unable to identify them as ciliates or naked amoebae.

*Needles penetrated by enchytraeids*
<figure><img src="image_15.png" /><figcaption>Fig. 91: Needle with a development of bacteria and micro-algae, section showing remains of transfusion tissue (CP); Fig. 92: Idem, section showing a micro-algal colony (Chlorophyceae) at the inside of a needle (CP); Fig. 93: Idem, section showing micro-algal colonies of Cyanophyceae (BMSL); Fig. 94: Idem, section showing protozoan cysts at the inside of a needle (BMSL); Fig. 95: Needle penetrated by enchytraeids, section showing mucus deposits, with micro-algae, along a gallery (BMSL); Fig. 96: Idem, section showing mucus deposits with bacteria only (BMSL); Fig. 97: Idem, section showing a pollen grain of pine at the inside of a gallery (CP); Fig. 98: Idem, section showing a thecamoeban cyst at the inside of a gallery (CP)</figcaption></figure>

CP = phase contrast; BMSL = methyl blue-Sudan III-lactophenol coloration; ar. = areolae; c.a. = micro-algal colonies; k.P. = protozoan cysts; a. = micro-algae; b. = bacteria; g.p. = pollen grains; th. = thecamoeban cysts; length of segment = 50 µm
In addition to numerous bites and breaks made by these animals, some needles were penetrated by these oligochetes, more often than in L₁ where such penetration was exceptional. Mucus was deposited along galleries bored at the inside of pine needles by enchytraeids, with a development of micro-algae (Fig. 95) or bacteria (Fig. 96). Other items transported by these animals (see observations on enchytraeid worms in L₁) were deposited within their galleries, such as pollen pine (Fig. 97) or cysts of small Thecamoeba such as *Euglypha* spp. (Fig. 98).

When enchytraeids deposited their faeces at the inside of decaying pine needles (which was not always the case) then the excreted plant material (strongly degraded by fungi then by enchytraeids) was no longer recognizable (Fig. 99). Hyphal fragments can be observed, without any cell content and thus neither stained by methyl blue nor showing any opacity under phase contrast (Frankland 1974, 1975). Faeces deposited by enchytraeids at the inside of pine needles were the seat of an intense bacterial development (Figs. 100 & 101).

Remaining tissues were in an advanced stage of decomposition, with bacteria, algae and protozoa (Figs. 102 & 103). Numerous brown spherules of varying size were observed in the mesophyll (Fig. 104). Their origin could be deduced from observations made in less degraded tissues such as metaxylem (Fig. 105): they were issued from the condensation of a brown substance which was deposited on cell walls of pine tissues. Similar brown deposits were described by authors who studied in microscopy the decomposition of beech and oak leaves. Olah et al. (1978) were speaking of brown substances originating from condensation and oxidation of cell cytoplasm. Toutain (1981) interpreted them as brown pigments originating from polyphenol-protein complexes formed from the reaction of tannins (contained in vacuoles in living cells) and proteins, which occurs at the death of cells. This author considered this process as being responsible for the brown colour of dead leaves. If the formation of such condensed products by chemical reaction between proteins and tanning substances was experimentally demonstrated by Handley (1954) and Kononova (1961), no proof can be given that the same process occurred in the pine material here described. In the present material, brown spherules were observed in metaxylem tracheids, i.e. in cells without any cytoplasm but which could be filled with sap at the time of needle fall. Anyway, brown spherules were not observed in all needles, even those in an advanced stage of decomposition, and they could not be taken responsible for the observed darkening of needles.

*Hyphal networks embedding decaying pine needles*

Loose mycelial wefts were observed surrounding pine needles, sometimes agglutinating them (Fig. 106). They were comprised of two different fungi: the secondary mycelium with clamp connections (3 µm hyphal diameter) of a hyaline basidiomycete which was identified as *Hyphodontia sp*. (Fig. 107), and melanised hyphae (diameter 4 µm) of an ascomycete which was identified as *Cenococcum geophilum* Fr. (Fig. 108). As will be seen below these two fungi could be identified through a panel of morphological characters during the study of mycorrhizae and decaying wood. Cell walls of basidiomycetous hyphae were often seen encrusted with oxalate crystals (Fig. 108).

Hyphal networks were inhabited by minute animals such as nematodes. Faeces of mesofauna were also found, the content of which revealed that the secondary mycelium of *Hyphodontia* was actively consumed (Fig. 109). In the faecal material, hyaline fungal hyphae were deprived of oxalate
<figure><img src="image_16.png" /><figcaption>Fig. 99: Needle penetrated by enchytraeids, section through enchytraeid faeces showing their content (CP); Fig. 100: Idem, with bacterial colonies (BMSL); Fig. 101: Idem, showing another type of bacterial colonies (BMSL); Fig. 102: Idem, section showing protozoan cysts at the inside of a needle (BMSL); Fig. 103: Idem, section through mesophyll showing micro-algae and bacteria (BMSL); Fig. 104: Idem, showing brown spherules (CP); Fig. 105: Idem, brown spherules in metaxylem (BMSL); Fig. 106: Hyphal network embedding decaying pine needles, overall view (CP)</figcaption></figure>

CP = phase contrast; BMSL = methyl blue-Sudan III-lactophenol coloration; f.h. = fungal hyphae; c.b. = bacterial colonies; k.P. = protozoan cysts; a. = micro-algae; s.b. = brown spherules; length of segment = 50 µm
crystals and had a fuzzy contour, indicating a degraded state. The absence of any bacteria indicated that their degradation could have occurred in animal guts.

### Conclusions about decaying pine needles

The degradation of internal tissues of pine needles through the development of *Verticicladium trifidum*, which was initiated in the L₁ layer, further developed in the L₂ layer. However, another saprophagous fungal species was observed (the basidiomycete *Marasmius androsaceus*), a secondary (or tertiary) fungal infection which probably took place during winter or spring. The lignin-free mesophyll completely disappeared under the cellulolytic action of these fungi, and the lignified transfusion tissue was degraded by *Verticicladium trifidum*. The latter species was considered as the main agent of needle blackening, a process taking place on the internal side of hypodermis.

Fauna, in particular enchytraeid worms, which were abundant at the time of sampling, opened needles by nibbling or biting the coarse layers (cuticle, epidermis, hypodermis) which isolated internal tissues from the external environment. Viable propagules of micro-algae and bacteria were carried on the tegument of these animals, and were propagated through their tunnelling activity at the inside of pine needles, thereby initiating a bacterial stage of needle degradation, which was accompanied by protozoan development. In some needles at an advanced stage of decomposition, the disappearance of internal tissues was probably completed within 8-9 months. Bacterial and micro-algal colonies developed both in internal tissues and in the numerous folds of collapsed needles. The shape of these colonies was reminiscent of what had been observed at greater depth in soil micro-aggregates (Arpin et al., 1980).

Decaying needles were covered by a network of melanised hyphae, mostly dead (probably relict from previous winter and spring), and were embedded in the oxalate-rich mycelium of the basidiomycete *Hyphodontia sp*.

### Decaying pine twigs

Compared with the L₁ layer, where only tissues with cellulosic cell walls were invaded and degraded by fungi imperfecti, bacterial development was visible in the L₂ layer, even at the inside of metaxylem tracheids (Fig. 110). Previous fungal development was attested by the presence of dead hyaline hyphae, which could not be stained by methyl blue and were perfectly transparent under phase contrast. Tracheid walls exhibited incipient degradation (Fig. 111). Since no basidiomycetous development was observed, the succession of organisms taking place in the decomposition of pine twigs was ascomycetes (imperfect stages) → bacteria.

Dead hyphae of dematiaceous fungi where retrieved in vascular rays (see Figure 34 for living stages found in L₁), and numerous micro-algal colonies were observed in bark, quite similar to those found in L₁ (Fig. 38). Mesofaunal faeces were deposited at the surface of decaying pine twigs: their content was made of bacteria and dead fragmented melanised hyphae (Fig. 112).
<figure><img src="image_17.png" /></figure>

Fig. 107: Hyphal network embedding decaying pine needles, detail showing the secondary mycelium of the basidiomycete *Hyphodontia* sp. (CP); Fig. 108: Idem, detail showing melanised hyphae of the dematiaceous ascomycete *Cenococcum geophilum* mixed with hyaline hyphae of *Hyphodontia* sp. (CP); Fig. 109: Idem, detail showing the content of embedded mesofaunal faeces (CP); Fig. 110: Pine twig, longitudinal section showing metaxylem tracheids, with empty hyphae and bacteria (CP); Fig. 111: Idem, showing degraded tracheid walls (CP); Fig. 112: Idem, showing the content of deposited mesofaunal faeces (BMSL); Fig. 113: Pine euphyll, tangential optical section through epidermis (LN); Fig. 114: Idem, tangential optical section showing a superficial network of melanised hyphae (LN)

CP = phase contrast; LN = natural light without coloration; a.a. = clamp connections; h.m. = melanised hyphae; p.i. = fungal cell walls encrusted with oxalate crystals; length of segment = 50 µm
Decaying euphylls of pine

Euphylls (flower and needle bracts) of pine were intact, which could be explained by their membranous texture and their recent fall (June). Euphyll epidermis exhibited a typical aspect (Fig. 113), quite different from that of pseudophylls (needles). They were covered by a network of poorly ramified melanised hyphae of 4 µm diameter (Fig. 114), quite similar to those attributed to *Cenococcum geophilum* in the mycelial network embedding needles (Fig. 108). Similarly to needles, pine euphylls were embedded in the mycelium weft formed by *Hyphodontia sp.* and *Cenococcum geophilum*.

Oribatid faeces were deposited at the surface of pine euphylls: their content was made of melanised fungal material and they were embedded in the mycelial network formed by *Hyphodontia sp.*

Decaying male flowers of pine

Bacteria, micro-algae and numerous quartz grains of silt size were observed at the inside of decaying male inflorescences of pine (Fig. 115). This association of silt-size mineral particles, bacteria and micro-algae was reminiscent of the material transported on the tegument of enchytraeids in the L₁ layer (Fig. 45), indicating that these worms penetrated male inflorescences, in which they fed on pollen (Fig. 51). The surface of quartz grains was punctuated with brown dots, which I interpreted as bacterial corpses.

Decaying branchwood of pine

The age of the young branch of pine which was collected in my sample was estimated to five years, taking into account the successive development of early- and latewood. From place to place bark was peeled off and sometimes was lacking. Below bark, external wood was softened to a great extent, being translucent and spotted black. Between bark and wood there was a fine layer of fine organic matter, with numerous enchytraeids, some of which were penetrating bark. This layer was made of enchytraeid faeces, which will be described below with the faunal material.

At the surface of wood, in those places where it was not covered by bark, there was a carpet of conidiophores of *Haplographium delicatum* Berck. & Br. (Fig. 117), which is the conidial (imperfect) stage of the ascomycete *Hyaloscypha dematiicola* (Berck. & Br.). Deeper in the wood, no prominent degradation was visible under the dissecting microscope, except that vessels were easily separated when crashed.
<figure><img src="image_18.png" /><figcaption>Fig. 115: Male flower of pine, detail showing quartz grains of silt size and bacteria (BMSL); Fig. 116: Idem, detail showing quartz grains with brown dots (BMSL); Fig. 117: Young pine branch, conidiophore of *Haplographium delicatum* (CP); Fig. 118: Idem, section through bark showing decaying cell walls and brown deposits (BMSL); Fig. 119: Idem, section showing bacterial colonies within bark (CP); Fig. 120: Idem, longitudinal section showing the secondary mycelium of *Hyphodontia sp.*, with cystidia, in metaxylem tracheids (CP); Fig. 121: Idem, showing chains of fungal spores and cystidia (CP); Fig. 122: Idem, showing degraded bordered pits (areolae) (CP)</figcaption></figure>

CP = phase contrast; BMSL = methyl blue-Sudan III-lactophenol coloration; g.q. = quartz grains; p.b. = brown dots; b.b. = brown deposits; c. = cystidia; sp. = spores; ar. = areolae; length of segment = 50 µm
## Bark

Bark was in a stage of degradation, contrary to what was observed in L₁. Brown deposits, which were described at the inside of peridermic cells in L₁ (Figs. 40, 41 & 42), remained intact and were covered with bacteria (Fig. 118). Numerous micro-sites showed bacterial colonies (Fig. 119).

## Wood

The internal part of wood was invaded by the secondary mycelium with clamp connections of *Hyphodontia* sp. (Fig. 120). Its hyphae were thinner (diameter 2 µm) and grew differently from what was observed in the mycelium embedding needles (Fig. 107). Numerous cystidia (short hyphae excreting oxalic acid) of particular size and shape were observed (Figs. 120 & 121), which were used for the identification of this fungus thanks to their taxonomic value at the species-group level. Cystidial types of the same kind have been described in cultures of wood-inhabiting basidiomycetes of the *Odontia* group (Corticiaceae) such as *Echinodontium tinctorium* (Ell. & Ev.) according to Nobles (1965), and *Grandinia farinacea* (Pers.) according to Käärik & Rennerfelt (1957). Based on the particular morphology of its cystidia (lagenocystidia), this fungus was considered as belonging to one of the three species *Hyphodontia arguta* (Fr.) Erikss., *Hyphodontia pallidula* (Bres.) Erikss. or *Hyphodontia alutaria* (Burt) Erikss. (Boidin, personal communication).

Tracheids with bordered pits appeared to be degraded in peripheral branchwood, areolae having enlarged and lost their torus (Fig. 122). In the wood located just under bark, numerous spores, isolated or in chains (Fig. 121), were found, belonging probably to the genus *Scytalidium* Pesante (fungi imperfecti). In the zones deprived of bark cover, hyaline hyphae, wide (diameter 7-10 µm) and slightly stained by methyl blue (Fig. 123), were followed and were seen to be connected to conidiophores of *Haplographium delicatum*, thereby allowing identification of the mycelium. In enchytraeid faeces deposited between bark and wood, I found hyaline hyphae still living at sampling time, growing apices of which being strongly stained by methyl blue (Fig. 124). The pattern of fungal colonisation observed in the peripheral zone of branchwood was interpreted as a succession from ascomycetes (fungi imperfecti) to basidiomycetes, the latter (*Hyphodontia*) being able to penetrate more deeply into the wood, and one of the ascomycetes continuing its growth in the mass of enchytraeid faeces accumulated between bark and wood. Such sharing of space and time by several fungal species colonizing the same pine branch was reminiscent of observations done by Butcher (1968) on *Pinus radiata* D. Don., i.e. superficial colonisation by several ascomycetes, followed a long time after by basidiomycetes penetrating deeper at the inside. More complicated patterns of succession were observed by Mangenot (1952) and Meredith (1959, 1960) when tree trunks were previously attacked by fungal parasites when still living. In their scheme first colonisers of fallen trunks were parasites, which continued their growth in dead wood, as I observed in some pine needles infected on the tree before decomposing in litter, then secondary and tertiary colonisers. If primary colonisation by parasites is skipped then a similar pattern emerges, with fungi imperfecti fructifying in places from which bark has been peeled off (Mangenot, 1952), followed by a wave of saprophagous basidiomycetes eliminating previous heartwood dwellers through antagonisms (Meredith, 1960).
<figure><img src="image_19.png" /></figure>

Fig. 123: Young pine branch, detail showing dead hyaline hyphae of *Haplographium delicatum* (BMSL); Fig. 124: Idem, detail showing intact hyaline hyphae with growing apices in enchytraeid faeces deposited between bark and wood (BMSL); Fig. 125: Idem, detail showing resin with crystals (LN); Fig. 126: Mycorrhiza, transversal section through fungal sheath and extra-matrical mycelium (CP); Fig. 127: Idem, detail of the extra-matrical mycelium, showing hyphae encrusted with oxalate crystals and smooth growing apices (BMSL); Fig. 128: Stem of the moss *Pseudoscleropodium purum*, longitudinal section through parenchyma, showing dead cells filled with micro-algae and bacteria (BMSL); Fig. 129: Leaf of the moss *Pseudoscleropodium purum*, showing intact cell walls (CP); Fig. 130: Idem, showing cells with bacterial development

CP = phase contrast; BMSL = methyl blue-Sudan III-lactophenol coloration; LN = natural light without coloration; h.h. = hyaline hyphae; ap. = apices; cr. = crystals; a.a. = clamp connections; p.i. = fungal cell walls encrusted with oxalate crystals; length of segment = 50 µm
It must be noticed, too, that resin was abundant in the peripheral zone of the studied branch, with rhombohedral crystals of unknown origin clearly visible in it (Fig. 125). Resin was accumulated between bark and wood, where it seemed to have been actively consumed by enchytraeids then retrieved in their faeces, as it will be described below.

## Pine ectomycorrhizae

The fine root system of pine seemed near entirely mycorrhizal in the studied sample. Fine roots were not very abundant in the L₂ layer, since observation of the humus profile to the naked eye revealed fine roots mostly in F₂ and H₁ (Table 1). However, they were far from negligible in this very superficial layer. According to Dominik (1969), most mycorrhizae found in the L₂ layer could be classified in the Ae type, i.e. ectotrophic mycorrhizae (ectomycorrhizae) with a diffuse woolly extra-matrical mycelium, a prosenchymatous fungal sheath and a Hartig net.

The extra-matrical mycelium was constituted by a network of hyaline hyphae with clamp connections, morphologically identical to those observed embedding needles and euphylls, except that their cell walls were smooth (Fig. 126). In more remote parts of the extra-matrical mycelium, hyphae appeared encrusted with oxalate crystals, only growing apices being smooth (Fig. 127).

At this stage of the present study, a question arose about the nature of the fungi which embedded most plant remains: were they mycorrhizal or not? The answer was given while studying the underlying F₁ layer, as will be explained below.

## Decaying moss (*Pseudoscleropodium purum*)

### Stems

Cells of moss stems were invaded by micro-algae and bacteria (Fig. 128), and cell walls seemed degraded. These observations did not cope with the study done by Kilbertus et al. (1970) on the same species and in quite similar environmental conditions. These authors described cells with intact walls, however according to their photographs it seems that they studied thick-walled cells belonging to a supporting tissue similar to collenchyma rather than parenchymatous cells with thin walls, like I did it.

### Leaves

When moss leaves were not torn internal attacks by microorganisms were never observed: cells were devoid of cytoplasm but walls were intact, middle lamella comprised (Fig. 129). When leaves were wounded, most probably by enchytraeids which abounded in moss material, then an intense bacterial
development (cocci) was observed to occur at the inside of broken cells, filling them near entirely (Fig. 130).

Most often moss leaves were embedded in the secondary mycelium of *Hyphodontia sp.*, which was frequently associated with that of another fungus, with clamp connections and smooth hyphae of a bigger (10 µm) diameter with thicker walls (Fig. 131).

Compared with the L₁ layer, it is worth to notice the disappearance of the sterile dematiaceous mycelia crawling at the surface of moss leaves, and its replacement by a hyaline woolly basidiomycetous mycelium, without any direct contact with the surface of leaves. These observations are in agreement with those of Kilbertus (1968a, b) and Kilbertus et al. (1970) on decaying *Pseudoscleropodium purum*. The white basidiomycete they described, hyphae of which were encrusted with oxalate crystals, is strongly reminiscent of the mycelium here described embedding mycorrhizae and permeating decaying litter. As in the present study, they thought it to be mycorrhizal. In addition, Kilbertus (1972) showed that there was a correlation between the abundance of *Pseudoscleropodium purum* and that of pine mycorrhizae in the top few centimetres, pointing on a possible influence of the moss on the process of mycorrhization, with the fungus as mediator.

Another question concerns the disappearance in L₂ of the dematiaceous mycelium which was observed at the surface of moss leaves in L₁ (Fig. 44). Given the amount of melanised hyphae which were found in faeces and guts of animals (oribatid mites, enchytraeids, fly larvae), it could be hypothesized that this fungus material was highly palatable to and was heavily grazed by small litter invertebrates. However, melanised hyphae were still visible at the surface of pine needles in L₂, although in a denser network where the visible impact of grazing was more limited than in L₁.

## Decaying bracken foliage (*Pteridium aquilinum*)

### *Leaflets*

Leaflets at several stages of decomposition were found, of which only two contrasted stages will be presented in the following.

* Leaflets with limb still present

To the exclusion of veins, only the upper epidermis was still present. Typical articulated epidermal hairs were observed (Fig. 132). Epidermal cells were invaded by hyaline hyphae (Fig. 133), as well as stomatal guard cells (Fig. 134). Similar to pine needles and moss shoots, bracken leaflets were embedded in the mixed mycelium formed by hyaline hyphae with clamp connections and oxalate crystals of the basidiomycete *Hyphodontia sp.* and by melanised hyphae of the ascomycete *Cenococcum geophilum* (Fig. 135). Previous fungal development (maybe before bracken fronds fell on the ground) was attested by the presence of empty fructifications (thyriothecia) of ascomycetous
<figure><img src="image_20.png" /></figure>

Fig. 131: Mycelium embedding leafy shoots of the moss *Pseudoscleropodium purum*, showing two kinds of basidiomycetous hyphae with clamp connections (CP); Fig. 132: Decaying leaflets of bracken, *Pteridium aquilinum*, detail showing an articulated epidermal hair (CP); Fig. 133: Idem, detail showing hyaline hyphae in epidermal cells (CP); Fig. 134: Idem, detail showing hyaline hyphae in stomatal guard cells (CP); Fig. 135: Idem, detail showing hyaline and melanised hyphae embedding foliar material (CP); Fig. 136: Idem, detail showing a thyriothecium of Microthyriaceae on the limb surface (CP); Fig. 137: Idem, detail showing a limb part nibbled by fauna, leaving intact a vein (CP); Fig. 138: Idem, detail showing a bacterial colony and fungal hyphae in epidermal cells (CP)

CP = phase contrast; p.e. = epidermal hairs; c.g. = guard cells; h.h. = hyaline hyphae; h.m. = melanised hyphae; p.i. = fungal cell walls with oxalate crystals; c.b. = bacterial colonies; length of segment = 50 µm
Microthyriaceae (Fig. 136). Faeces of oribatid mites were also observed, containing only fungal material.

* Vein networks

At this stage of bracken leaflet degradation, only networks of veins were still visible. Traces of nibbling by small invertebrates were present on both sides of remaining veins (Fig. 137). Enchytraeids were thought responsible for the disappearance of bracken limbs, given that their round-shaped, fuzzy-contoured faeces were found agglomerating at the surface of leaflets, their content asserting they had consumed bracken litter (see below).

Some small areas of upper epidermis, which were still present at this advanced stage of degradation, showed a bacterial development at the inside of epidermal cells (Fig. 138). Fungal loculae (pycnidia, ascocarps?) were found still adhering at the surface of intact veins or included in the dense fungal mycelium which replaced degraded bracken material (Fig. 139).

*Rachises*

The material present in the L₂ sample corresponded to secondary or tertiary rachises of *Pteridium aquilinum*, and not to primary rachises (bracken petioles). On the same fragment, several stages of decomposition could be found in the parenchymatous tissue. In some places, cell walls were still intact but the cytoplasm was invaded by the secondary mycelium with smooth walls of an unidentified basidiomycete (Fig. 140). In other places parenchymatous cells were degraded and invaded by rod-shaped bacteria (Fig. 141) or cocci (Fig. 142). Scalariform tracheids remained intact, even though they were invaded by hyphae of the same unidentified basidiomycete (Fig. 143). The examination of enchytraeid faeces which had been deposited at the surface of a bark fragment revealed the same bracken tracheids (Fig. 144) thereby indicating that enchytraeids consumed internal tissues of bracken rachises.

Comparison with results of other authors was difficult, given the scarcity of studies dealing with the decomposition of bracken foliage. Frankland (1966, 1969) studied fungal successions over two years of decomposition of bracken petioles and she showed that fauna intervened occasionally but not before the second year. In the present study I observed a much more rapid development of the decay process, the passage to a bacterial stage and concomitant ingestion by fauna (enchytraeids) occurring around six months after the fall of dead foliage. However, it must be noticed that the bracken frond ramifications I studied were thinner than petioles.

Fauna
<figure><img src="image_21.png" /><figcaption>Fig. 139: Decaying leaflets of bracken, *Pteridium aquilinum*, detail showing thyriothecia of Microthyriaceae embedded in the secondary mycelium of *Hyphodontia sp*. (CP); Fig. 140: Decaying rachis of bracken, *Pteridium aquilinum*, longitudinal section showing parenchymatous cells with hyaline hyphae (BMSL); Fig. 141: Idem, detail showing rod-shaped bacteria (CP); Fig. 142: Idem, detail showing coccoid bacteria (BMSL); Fig. 143: Idem, detail showing a scalariform tracheid with hyaline hyphae at the inside (BMSL); Fig. 144: Similar scalariform tracheid in an enchytraeid excrement, with a melanised hypha invaded by bacteria (BMSL); Fig. 145: Enchytraeid worm observed by transparency, showing the zigzag shape of its empty intestine (CP); Fig. 146: Idem, overall view of gut contents observed by transparency (LN)</figcaption></figure>

CP = phase contrast; BMSL = methyl blue-Sudan III-lactophenol coloration; LN = natural light without coloration; h.h. = hyaline hyphae; a.a. = clamp connections; b. = bacteria; length of segment = 50 µm
Table 5 shows biovolumes and densities of all animal groups which were found during sorting of L₂ litter. Compared to L₁, the diversity of animal groups was slightly greater, and oribatid mites decreased while enchytraeids increased in abundance. While the total biovolume of oribatids decreased by a factor of 10, that of enchytraeids increased by the same factor. Some care is requested when comparing data for springtails (Collembola), because a single specimen of *Pogonognathellus flavescens* occupied the totality of the biovolume of springtails in L₁, while several small species were present in L₂. Among them, two isotomids displayed contrasted distributional patterns between L₁ and L₂: *Parisotoma notabilis*, which was dominant in numbers in L₁, was replaced by *Folsomia manolachei* Bagnall. Fly larvae remained approximately at the same level of biovolume, but Cecidomyidae, dominant in L₁, were replaced by Sciaridae, and nymphs of these insects appeared in L₂. Several groups of tiny invertebrates were represented by single specimens: tardigrades, pauropods and copepods.

<table><caption>**Table 5.** Taxonomic name, abundance and estimated biovolume of living animals found in the L₂ layer</caption><thead><tr><th>Taxonomic groups and species</th><th>Abundance</th><th>Upper estimate of biovolume (µL)</th><th>Lower estimate of biovolume (µL)</th></tr></thead><tbody><tr><td>Nematodes</td><td>21</td><td>0.10 10<sup>-1</sup></td><td>0.03 10<sup>-3</sup></td></tr><tr><td>Dorylaimidae</td><td>8</td><td>0.60 10<sup>-2</sup></td><td>0.15 10<sup>-2</sup></td></tr><tr><td>Anaplectus sp.</td><td>6</td><td>0.16 10<sup>-2</sup></td><td>0.04 10<sup>-2</sup></td></tr><tr><td>Tylolaimophorus sp.</td><td>4</td><td>0.63 10<sup>-3</sup></td><td>0.16 10<sup>-3</sup></td></tr><tr><td>Prionchulus sp.</td><td>2</td><td>0.18 10<sup>-2</sup></td><td>0.05 10<sup>-2</sup></td></tr><tr><td>Clarkus papillatus</td><td>1</td><td>0.25 10<sup>-3</sup></td><td>0.06 10<sup>-3</sup></td></tr><tr><td>Enchytraeids</td><td>39</td><td>2.6</td><td>0.7</td></tr><tr><td>Tardigrades</td><td>1</td><td>0.95 10<sup>-3</sup></td><td>0.24 10<sup>-3</sup></td></tr><tr><td>Pauropods</td><td>1</td><td>0.27 10<sup>-3</sup></td><td>0.07 10<sup>-3</sup></td></tr><tr><td>Oribatid mites</td><td>12</td><td>0.92 10<sup>-1</sup></td><td>0.23 10<sup>-1</sup></td></tr><tr><td>Oppia sp.</td><td>4</td><td>0.81 10<sup>-2</sup></td><td>0.20 10<sup>-2</sup></td></tr><tr><td>Platynothrus peltifer</td><td>3</td><td>0.60 10<sup>-1</sup></td><td>0.15 10<sup>-1</sup></td></tr><tr><td>Nanhermannia nana</td><td>2</td><td>0.29 10<sup>-2</sup></td><td>0.07 10<sup>-2</sup></td></tr><tr><td>Adoristes ovatus</td><td>1</td><td>0.16 10<sup>-1</sup></td><td>0.04 10<sup>-1</sup></td></tr><tr><td>Suctobelba sp.</td><td>1</td><td>0.97 10<sup>-3</sup></td><td>0.24 10<sup>-3</sup></td></tr><tr><td>Belboidea sp.</td><td>1</td><td>0.48 10<sup>-2</sup></td><td>0.12 10<sup>-2</sup></td></tr><tr><td>Miscellaneous mites</td><td>5</td><td>0.42 10<sup>-1</sup></td><td>0.11 10<sup>-1</sup></td></tr><tr><td>Gamasina</td><td>2</td><td>0.34 10<sup>-1</sup></td><td>0.09 10<sup>-1</sup></td></tr><tr><td>Uropodina</td><td>1</td><td>0.59 10<sup>-2</sup></td><td>0.15 10<sup>-2</sup></td></tr><tr><td>Tydeidae</td><td>1</td><td>0.47 10<sup>-3</sup></td><td>0.12 10<sup>-3</sup></td></tr><tr><td>Acaridida</td><td>1</td><td>0.17 10<sup>-2</sup></td><td>0.04 10<sup>-2</sup></td></tr><tr><td>Copepods</td><td>1</td><td>0.95 10<sup>-3</sup></td><td>0.24 10<sup>-3</sup></td></tr><tr><td>Springtails</td><td>12</td><td>0.18 10<sup>-1</sup></td><td>0.04 10<sup>-1</sup></td></tr><tr><td>Folsomia manolachei</td><td>7</td><td>0.10 10<sup>-1</sup></td><td>0.03 10<sup>-1</sup></td></tr><tr><td>Isotomiella minor</td><td>1</td><td>0.17 10<sup>-2</sup></td><td>0.04 10<sup>-2</sup></td></tr><tr><td>Allacma fusca</td><td>1</td><td>0.20 10<sup>-2</sup></td><td>0.05 10<sup>-2</sup></td></tr><tr><td>Lepidocytus lanuginosus</td><td>1</td><td>0.10 10<sup>-2</sup></td><td>0.03 10<sup>-2</sup></td></tr><tr><td>Parisotoma notabilis</td><td>1</td><td>0.11 10<sup>-2</sup></td><td>0.03 10<sup>-2</sup></td></tr><tr><td>Proisotoma minima</td><td>1</td><td>0.20 10<sup>-2</sup></td><td>0.05 10<sup>-2</sup></td></tr><tr><td>Fly larvae</td><td>12</td><td>0.33</td><td>0.08</td></tr><tr><td>Sciaridae</td><td>8</td><td>0.33</td><td>0.08</td></tr><tr><td>Cecidomyidae</td><td>3</td><td>0.22 10<sup>-2</sup></td><td>0.06 10<sup>-2</sup></td></tr><tr><td>Chironomidae</td><td>1</td><td>0.61 10<sup>-2</sup></td><td>0.15 10<sup>-2</sup></td></tr><tr><td>Fly nymphs</td><td>2</td><td>0.25</td><td>0.06</td></tr><tr><td>Sciaridae</td><td>2</td><td>0.25</td><td>0.06</td></tr></tbody></table>
Concerning microfauna, as this has been highlighted above, it should be clear that the present study can give only a truncated view of their importance in the studied litter profile. Despite of these shortcomings, numerous observations were made on cyst amoebae, other protozoa and nematodes. However, given the number of specimens of nematodes collected in L₂ (21) and the easiness with which they could be collected when seen under the dissecting microscope, this group was taken into account in Table 5. The fact that only one specimen of this group was observed in L₁, while 21 were observed with the same method and care in L₂, was indicative of their clear preference for the latter layer.

## Enchytraeid worms

Generally speaking the intestines of enchytraeids showed a great variety of contents when observed by transparency in chloral-lactophenol. Their zigzag shape (Figs. 145 & 146) was reminiscent of the regular subdivision of the body into numerous metameres and explained why their faeces were loosely assembled in feeding places where they were deposited. Sometimes, but rarely, only one type of gut content was observed filling the intestine, such as for instance resin (Fig. 147). More generally, gut contents were composite, without any clear dominance of a category. When fungal material was particularly abundant; it was often accompanied by plant fragments (tracheids, for instance) which indicated that the animal had ingested internal tissues of needles or wood, together with their fungal colonisers. The most prominent character of enchytraeid gut contents, compared to other invertebrates, was the apparent absence of selective feeding.

Most hyaline hyphae ingested by enchytraeids corresponded to the secondary mycelium of the basidiomycete *Hyphodontia sp.* which embedded litter debris. These hyphae were often in way to be degraded in enchytraeid intestines (Figs. 148 & 149). Some hyaline hyphae of another type were also observed: their thicker walls did not appear degraded (Fig. 150).

Melanised hyphae of *Cenococcum geophilum* showed intact walls, the contrast with hyaline hyphae being better visible when both items were present in close vicinity (Fig. 151). Some melanised fungal walls were perforated (Fig. 152), resembling figures described for the attack of fungal walls by bacteria (Olah et al., 1978). In this case, it is highly probable that hyphae were in this state of degradation before being ingested. In some other cases melanised hyphae appeared to be more seriously degraded, their walls being made thinner and softer (Fig. 153). This could be due to the action of the animal, however, it may be questioned whether enchytraeids were capable to degrade fungal walls impregnated with melanin at first ingestion. Reisinger (1972, 1978) showed that at least two successive ingestions by the mite *Tyrophagus putrescentiae*, with microbial development in the meantime, were necessary to degrade melanised fungal walls. Other observations showed that some kind of melanised fungal material could be directly degraded by enchytraeids. Several individuals were shown to transport big dematiaceous (internally echinulate) spores of *Sphaeropsis sapinea* (Fr.) (Fig. 154). These spores were retrieved in the intestine, with their walls at several stages of degradation (Figs. 155 & 156). Thus enchytraeids, at least the species here studied, which I presumed to be *Cognettia sphagnetorum* (Vejdovský), had the potential to degrade melanised fungal walls as soon as first ingestion. However, the bulk of observations done on both guts and faeces showed that
<figure><img src="image_22.png" /><figcaption>Fig. 147: Enchytraeid worm observed by transparency, gut filled with resin with crystals (CP); Fig. 148: Idem, detail of gut contents showing hyaline and bacteria in way to be degraded (CP); Fig. 149: Idem, basidiomycetous hyaline hyphae in way to be degraded (CP); Fig. 150: Idem, hyaline hyphae with intact walls (CP); Fig. 151: Idem, hyaline hyphae in way to be degraded, melanised hypha with near intact walls (CP); Fig. 152: Idem, melanised hypha with numerous perforations indicated by arrows (CP); Fig. 153: Idem, melanised hyphae in way to be degraded (CP); Fig. 154: Spores of *Sphaeropsis sapinea* on the tegument of an enchytraeid worm (CP)</figcaption></figure>

CP = phase contrast; b. = bacteria; r.d.h.h. = dense networks of hyaline hyphae; a.a. = clamp connections; h.m. = melanised hyphae; sp. = spores; length of segment = 50 µm
<figure><img src="image_23.png" /><figcaption>Fig. 155: Spores of *Sphaeropsis sapinea* at the inside of an enchytraeid intestine observed by transparency (CP); Fig. 156: Idem, showing several stages of degradation of spore walls (CP); Fig. 157: Enchytraeid gut contents observed by transparency, detail showing intact unicellular micro-algae (CP); Fig. 158: Idem, filamentous blue-green alga (cyanobacteria) with intact walls (CP); Fig. 159: Idem, unicellular micro-algae at different stages of degradation under the action of intestinal microflora (CP); Fig. 160: Idem, filamentous blue-green alga with dissociated cells indicated by arrows (CP); Fig. 161: Idem, reticulated tracheid and plant cells in way to be degraded (CP); Fig. 162: Idem, metaxylem tracheid in way to be degraded (CP)</figcaption></figure>

CP = phase contrast; a. = algae; b. = bacteria; tr. = tracheid; length of segment = 50 µm
this was far from being realized being realized during the time of transit through the intestine, in
particular concerning hyphae of *Cenococcum geophilum*.

Algae were commonly observed in enchytraeid intestines, for instance unicellular
Chlorophyceae (Fig. 157), quite similar to those found proliferating in oribatid faeces in L₁ (Fig. 71),
or filamentous blue-green algae (Cyanophyceae), now called cyanobacteria (Fig. 158). In the first
case, algal cells were often seen to be viable: they were turgescent, with their cytoplasm exhibiting
opacity in phase contrast. Such features were never observed with fungal material, only walls of which
appeared intact, at best. Algal cells were observed to be attacked by some bacteria from intestinal
microflora: bacteria were seen agglutinated around algal cells, which became collapsed and devoid of
cytoplasm. Several stages of algal degradation are visible on Figure 159. Filamentous blue-green algae
(cyanobacteria) exhibited also some signs of degradation: cells were emptied and cells of the same file
were separated, keeping intact their rigid cellulosic wall (Fig. 160). All these figures indicated that
algae were somewhat hard to become digested, compared to fungal material which was more easily
degraded. Studies in transmission electron microscopy by Kilbertus & Vannier (1979) showed that
algae may pass intact through collembolan intestines.

Enchytraeid worms ingested needle plant tissues. Cells were always comminuted (Fig. 161).
Recognizable cells were only those of protoxylem (Fig. 161) and metaxylem (Fig. 162), although they
exhibited severe deformations. It is highly probable that the degradation of plant cell walls, which had
been performed in a first stage by fungi and bacteria, reached a further stage in enchytraeid intestines.
Resin, which abounds between bark and wood in fallen branches, was ingested, too. Crystals were
found included in the ingested resin, which was found either as an isolated piece (Fig. 163) or
completely filling the intestine (Fig. 147). Doubts may be raised about the nutritional value of pine
resin. Maybe it provided the worms with some oligo-elements? The abundance of this food item in
enchytraeid guts, often seen filling the whole intestine, may indicate it to be a preferred food resource.
Another argument is that, compared to dead moss material which was dominant in the environment
but was neglected by these animals in L₂ (but see above for living moss in L₁), resin was scarcely
represented in the environment while voraciously consumed.

Another item which was frequently consumed by enchytraeids in L₂ was invertebrate faeces,
most probably of microarthropods according to the small size of ingested masses (Fig. 164). These
faeces exhibited numerous bacteria which were in way to be degraded (fuzzy contours in phase
contrast) in enchytraeid intestines. Fragments of melanised hyphae, in an advanced stage of
degradation, were also observed in ingested faeces.

Four other components of the enchytraeid food regime must be noted, too, because their
abundance was far from negligible although they were not so much frequently encountered as the
abovementioned items. Those were cysts of Thecamoeba, such as for instance *Trinema* spp., always in
way to be degraded (Fig. 165), corpses or exuviae of microarthropods of which only setae were still
intact (Fig. 166), quartz grains (Fig. 167) and pollen grains, most often of pine (Fig. 168) but also of
other plant species growing in the study site such as the grass *Molinia caerulea* (L.) Moench (Fig.
169).

Intestinal microflora was richer and more diverse than in any other enchytraeid worm found in
L₁. It was observed in all individuals I scrutinized, and it was distributed in all intestinal metameres. In
addition to the minute bacteria already photographed surrounding algae (Fig. 159), swarms of slightly
curved rod-shaped bacteria (Fig. 170) and cocci (Fig. 171) were observed. Several bacterial types
<figure><img src="image_24.png" /></figure>

Fig. 163: Enchytraeid gut contents observed by transparency, detail showing a piece of resin with crystals (CP); Fig. 164: Idem, microarthropod faeces (CP); Fig. 165: Idem, cyst of *Trinema* sp. (cyst amoeba) in way to be degraded (CP); Fig. 166: Idem, microarthropod corpse or exuvia, in way to be degraded, and fungal spores (CP); Fig. 167: Idem, composite gut contents with quartz grains (CP); Fig. 168: Idem, fungal material, bacteria, fragment of pine pollen (CP); Fig. 169: Idem, pollen grain of *Molinia caerulea* with torn exine, fungal hyphae and bacteria in way to be degraded (CP); Fig. 170: Idem, intestinal microflora in the absence of food bolus (CP)

CP = phase contrast; cr. = crystal; f.h. = fragments of hyphae; ec. = scales; s. = setae; g.q. = quartz grains; g.p. = pollen grains; length of segment = 50 µm
could be present in the same intestine, but never in the same metamere. Direct contact of intestinal microflora with the food bolus was exceptional (Fig. 159). Customarily, bacteria of intestinal microflora were seen as swarms surrounding the food bolus, or dispersed in the intestinal lumen when devoid of food, as photographed here.

At first sight the observed enchytraeids did not seem to select food resources. However, the absence of moss material and the abundance of resin let us think that they exhibited preferences. Such a phenomenon has been evidenced in Collembola by McMillan (1975).

## Oribatid mites

Oribatid mites, which were the dominant invertebrate group in L₁, were replaced by enchytraeids in L₂. In this layer, dead oribatids (10 specimens) were near as abundant as living individuals (12 specimens), indicating that L₂ had been abandoned to the benefit of more superficial litter which was made of living moss and freshly fallen needles. At the inside of the oribatid group, it must be noted that big-sized species regressed in number, to the exception of *Nanhermannia nana* (Nicolet) which was absent from L₁ (Table 5), but small-sized species (mainly *Oppia* spp.) remained at the same level of abundance. Reasons for this apparent shift of oribatid species to more superficial litter were unclear, given that sampling took place during a dry and warm period (late August). Three non-conflicting hypotheses were raised to explain it, (i) intolerance of oribatid mites to the intense activity of enchytraeids in L₂, forcing them to live in the upper horizon L₁ where more drought-sensitive enchytraeids were at their disadvantage, (ii) intolerance to growing basidiomycetous mycelia (maybe causing a physical obstacle to foraging) or (iii) food preferences. Observations on the food bolus of oribatid mites seemed to sustain the third hypothesis, although the low number of non-starving individuals did not allow me to conclude neatly on that point.

*Platynothrus peltifer* showed the same intestinal bacteria as in L₁ (Fig. 61), in swarms in close proximity to food boluses which were made of strongly transformed fungal material (Fig. 172). Other views showed even more strongly decayed fungal material where only some hyphal fragments were still recognizable (Fig. 173). In all cases, bacterial development was noticeable. By comparison with L₁, it can be hypothesized that *Platynothrus peltifer* ingested faeces in L₂, most probably of oribatids given the small size of hyphal fragments found in them. Coprophagy had not been suspected in L₁.

All individuals of the genus *Oppia* which were not starving showed the same gut contents (Fig. 174): fragments of hyphae of *Hyphodontia* sp., with similar hyphal diameter and oxalate crystals as in the mycelium commonly embedding litter debris in L₂. Several stages of degradation of fungal walls were observed to occur at the inside of the intestine: first oxalate crystals were lost, and then fungal walls took fuzzy contours. The same basidiomycetous mycelium was never seen to be consumed by *Platynothrus peltifer*. My hypothesis was that *Platynothrus peltifer*, contrary to *Oppia* spp., did not find its preferred food in L₂ and rather ingested its own faeces which had been deposited some time before when L₂ was still a L₁ layer. Although numerous studies were performed on food habits of *Platynothrus peltifer* (Hartenstein 1962a, b; Littlewood, 1969; Pande & Berthet, 1973; Anderson, 1975; Behan-Pelletier & Hill, 1983), these authors did not determine the stage of degradation of the ingested material, thus no definite conclusion can be reached about food preferences of this species.
<figure><img src="image_25.png" /></figure>

Fig. 171: Enchytraeid gut contents observed by transparency, intestinal microflora in the absence of food bolus (CP); Fig. 172: Gut contents of the oribatid mite *Platynothrus peltifer* observed after dissection, detail showing intestinal microflora (right side) and food bolus (left side) made of decaying fungal material (CP); Fig. 173: Idem, detail showing strongly transformed fungal material and bacteria (CP); Fig. 174: Gut contents of the oribatid mite *Oppia* sp. observed after dissection, detail showing hyphae of the basidiomycete *Hyphodontia* sp. at different stages of degradation of cell walls and disappearance of oxalate crystals (CP); Fig. 175: Gut contents of the isotomid collembolan *Folsomia manolachei* observed by transparency, detail showing strongly decayed melanised fungal material and bacteria (CP); Fig. 176: Idem, detail showing strongly decayed and compacted melanised fungal spores (CP); Fig. 177: Gut contents of sciarid fly larvae observed by transparency, detail showing intestinal microflora (CP); Fig. 178: Idem, detail showing hyaline and melanised fungal hyphae, with intact walls (CP)

CP = phase contrast; f.h. = fragmented fungal hyphae; p.i. = fungal cell walls encrusted with oxalate crystals; sp. = spores; h.h. = hyaline hyphae; h.m. = melanised hyphae; length of segment = 50 µm
Springtails

Contrary to oribatid mites, Collembola were more numerous and species-rich in L₂ (12 specimens, 6 species) than in L₁ (9 specimens, 2 species). As in L₁, Isotomidae were dominant, *Parisotoma notabilis* being nevertheless replaced by *Folsomia manolachei*. As was observed in L₁, gut contents of isotomid springtails were mostly made of strongly degraded fungal material and bacteria, several stages of degradation being observed (Figs. 175 & 176). The latter photograph, displaying dematiaceous fungal spores compacted and in an advanced stage of transformation, is strongly reminiscent of what has been described as ‘humus’ or ‘amorphous organic matter’ by several authors (Knight & Angel, 1967; Bödvarsson, 1970; Gilmore & Raffensperger, 1970; McMillan & Healey, 1971; Hågvar & Kjøndal, 1981). My hypothesis, based on the great abundance of bacteria included in the food bolus, is that animal faeces were ingested, more particularly those of oribatid mites given the fungal nature of the food bolus and the small size of hyphal fragments. This was confirmed, as will be seen below, by the study of animal faeces found in L₂. Intestinal microflora was never observed in all collembolan specimens studied.

Fly larvae

The abundance of fly larvae was higher in L₂ (12 specimens) than in L₁ (7 specimens), but Cecidomyidae were replaced by Sciaridae, nymphs of which were also seen, in addition to larvae. Chironomidae remained at a very low level of abundance, as in L₁.

Sciarid larvae possessed an intestinal microflora (Fig. 177). Contrary to what had been observed in L₁ (Figs. 65 & 66), only fungal material was ingested in L₂. As for enchytraeids, dead moss material, cells of which were devoid of cytoplasm, was totally neglected. Fungal hyphae kept their walls intact, whether melanised (Fig. 178) or hyaline such as those of *Hyphodontia sp*. (Fig. 179). The absence of any degradation of hyaline basidiomycetous hyphae in sciarid intestines was exceptional and was not registered in any other invertebrate group I studied in this sample. Plant material was never observed in sciarid guts.

Cecidomyidae larvae, as was observed in L₁, did not possess any solid matter in their digestive tract. Liquid food can be suspected, as was hypothesized by Healey & Russel-Smith (1971) on the base of the anatomy of mouth pieces. However these animals transported numerous propagules on their tegument, probably viable according to phase contrast, for instance bacteria (Fig. 180) and microalgae (Fig. 181). The transport of microbial propagules adhering on a sticky (probably mucous) tegument is a character cecidomyid larvae had in common with enchytraeids.

Gut contents of Chironomidae were made of strongly degraded fungal material, mixed with numerous bacteria (Fig. 182), as this had been observed in isotomid springtails (Figs. 175 & 176). Comparing these results with the observations made in L₁ on larval chironomid guts (Fig. 70) led me to conclude that they ingested animal faeces. However, the direct ingestion of fungal hyphae had been
<figure><img src="image_26.png" /></figure>

Fig. 179: Gut contents of sciarid fly larvae observed by transparency, detail showing hyphae of *Hyphodontia sp.*, with intact walls (CP); Fig. 180: Bacteria transported on the tegument of cecidomyid fly larvae (CP); Fig. 181: Archethallus of Chlorophyceae transported on the tegument of cecidomyid fly larvae (CP); Fig. 182: Gut contents of chironomid fly larvae observed by transparency, detail showing strongly decayed fungal material and a melanised hypha with intact walls (CP); Fig. 183: Gut contents of the nematode *Anaplectus* sp. observed by transparency, showing intestinal microflora and small melanised fungal debris (CP); Fig. 184: Gut contents of the nematode *Tylolaimophorus* sp., showing intestinal microflora (CP); Fig. 185: The cyst amoeba *Nebela collaris* on a bracken leaflet of *Pteridium aquilinum* (CP); Fig. 186: Section through a macroinvertebrate excrement, detail showing a fragment of pine needle epidermis with a network of melanised hyphae (LN)

CP = phase contrast; LN = natural light without coloration; p.i. = fungal cell walls encrusted with oxalate crystals; a.a. = clamp connections; h.m. = melanised hyphae; f.h. = fragmented fungal hyphae; length of segment = 50 µm
also observed in L₁ (Fig. 69). Strong differences between sciarid and chironomid larvae both in food regime and impact on fungal walls have been also registered by Healey & Russel-Smith (1971).

### Nematodes

Nematodes seem to abound in L₂, at least when compared with L₁ where only one specimen of Monochidae had been collected. A few specimens showed gut contents, if we except very small debris of melanised fungal walls in a female *Anaplectus sp.* (Fig. 183), two mastax of rotifers (not shown, but see Fig. 58) in a larva of *Clarkus papillatus* (Bastian) and a bacterial mass in way to be degraded (not shown, but see Figs. 58-60) in a larva of *Dorylaimus* sp. However, intestinal microflora was always visible, applied to the intestinal epithelium, in *Anaplectus sp.* (Fig. 183) and *Tylolaimophorus sp.* (Fig. 184).

### Cyst amoebae

Cyst amoebae (Thecamoeba) were observed at the surface of plant material, mostly fern leaflets, when mounted for direct observation in chloral-lactophenol. Needles, which were sectioned for observing internal tissues, did not display cyst amoebae at the inside, but this failure can be ascribed to a methodological bias, as for most microfauna. Given that some sections showed cyst amoebae at the surface of collapsed pine needles (Fig. 94) it can be suspected that these protozoa abounded at the inside of decaying pine material, too. I observed *Nebela collaris* (Fig. 185), numerous *Euglypha sp.*, *Assulina semilunum* (Ehrenberg), *Phryganella acropodia* (Hertwig & Lesser) and *Corythion dubium* Taranek. It should be noticed that not any cyst amoebae had been found in the L₁ layer, to the exception of a specimen of *Nebela collaris* which had been ingested by an enchytraeid (Fig. 55), which indicates, if necessary, that L₂ was preferred to L₁.

### Animal faeces

#### Macroinvertebrate faeces

Some macroinvertebrate faeces had a convoluted contour which was reminiscent of earthworm faeces. However, most of them were tightly appressed against moss leafy shoots or pine needles and their shape was hardly recognizable. In this case they could be allocated as well to millipedes or woodlice. In the absence of clear faunal connection, and waiting for more material found underneath (but see F₁ layer), macroinvertebrate faeces collected in L₂ were not attributed to any animal group. However, it should be noticed that microscopic characters did not differ between them: all of them were comprised of coarse fragments of moss, pine and bracken plant tissues, which were poorly transformed, pointing
to a common zoological origin. Plant cells were in the same state as described in the study of non-
ingested decaying plant material and did not seem to be further degraded, for instance epidermis (Figs.
186 & 187), metaxylem tracheids (Fig. 188) or pine pollen grains (Fig. 189). However, some stomatal
guard cells were found isolated from the rest of the epidermis (Fig. 190). Other plant tissues have
disappeared, probably under the influence of the above described fungal development (as traced from
the presence of empty hyphae) and of an intense development of bacterial and cyanobacterial colonies
(Figs. 191 & 192). These microbial colonies did not differ from what has been observed in decaying
plant material, but they seemed to proliferate in macroinvertebrate faeces. Moss material was mainly
present in the form of entire leaves (Fig. 193) but sometimes moss cell walls were in way to be
degraded (Figs. 194 & 195), a pattern which had not been observed in non-ingested moss material.
Food selection did not seem evident, since not any plant or microbial material observable in L₁ and L₂
was absent from studied macroinvertebrate faeces. However, the particular abundance of corpses
and/or exuviae of arthropods should be noticed (Fig. 189), which cannot be explained by random
foraging. Living nematodes were observed in these faeces, as well as mycelia of the basidiomycete
*Hyphodontia sp.*

Near all macroinvertebrate faces were of a dark brown colour, due to their pure organic
content comprising empty hyphae of dematiaceous fungi. However, a unique specimen of faeces, of
convoluted contour and light colour, was found. Its content was purely mineral, made of quartz
particles of silt size without any bacterial development (Fig. 196). This was the first clue to mineral
uplift encountered in the studied profile, to the exception of mineral particles of similar nature which
were transported on the tegument of enchytraeid worms.

*Enchytraeid faeces*

Enchytraeid faeces were always observed as fluffy clusters (Fig. 197). Their composition differed
according to the place where they were deposited. Only plant tissues issued from needle consumption
were found in enchytraeid faeces at the inside of pine needles, as well as in faecal masses accumulated
between twin needles, for instance fragments (still intact) of hypodermis (Fig. 198) or stomatal guard
cells isolated among strongly degraded remains of needle tissues (Fig. 199). Plant material was always
mixed with numerous fragments of empty fungal hyphae. Fragments of pine tracheids and resin were
found in enchytraeid faeces accumulated between bark and wood, together with brown spores of
*Scytalidium* sp. (see Fig. 121). The mycelium of *Haplographium delicatum*, seen sporulating at the
surface of naked wood (Fig. 117) developed itself within fluffy masses of enchytraeid faeces
accumulated between bark and wood. Free clusters (not embedded in plant tissues) displayed
fragments of varied origin such as pine needles, wood, bark or bracken fern foliage (Fig. 144), but
never moss tissues. Faecal clusters were permeated by the mycelium of *Hyphodontia* sp., in direct
contact with decaying plant tissues (Fig. 200). In every case I noted an intense development of bacteria
(Fig. 201) and cyanobacteria (Figs. 202 & 203).

*Oribatid faeces*
<figure><img src="image_27.png" /></figure>

Fig. 187: Section through a macroinvertebrate excrement, detail showing a fragment of pine needle hypodermis, observed on its internal face (LN); Fig. 188: Idem, detail showing metaxylem tracheids of pine (CP); Fig. 189: Idem, detail showing a mite exuvia or corpse and a pine pollen grain (LN); Fig. 190: Idem, detail showing an isolated stomatal guard cell of pine (CP); Fig. 191: Idem, detail showing bacterial colonies (BMSL); Fig. 192: Idem, detail showing cyanobacterial colonies (BMSL); Fig. 193: Idem, detail showing a leaf of the moss *Pseudoscleropodium purum*, with intact walls (CP); Fig. 194: Idem, detail showing a fragment of moss leaf, with cell walls in way to be degraded (CP)

CP = phase contrast; LN = natural light without coloration; ar. = areolae; g.p. = pollen grains; c.b. = bacterial colonies; c.a. = cyanobacterial colonies; length of segment = 50 µm
<figure><img src="image_28.png" /></figure>

Fig. 195: Section through a macroinvertebrate excrement, detail showing a fragment of moss leaf, with cell walls in an advanced stage of degradation (CP); Fig. 196: Detail of macroinvertebrate excrement entirely comprised of silt-size quartz particles (CP); Fig. 197: Enchytraeid faeces clustered in a pine needle folding, overview (LN); Fig. 198: Idem, detail showing a fragment of pine needle hypodermis (CP); Fig. 199: Idem, detail showing a stomatal guard cell of pine needle (CP); Fig. 200: Enchytraeid faeces, detail showing pine tracheids with bordered pits and mycelium of *Hyphodontia sp.* (CP); Fig. 201: Idem, detail showing bacterial colonies (CP); Fig. 202: Idem, detail showing colonies of cyanobacteria (LN)

CP = phase contrast; LN = natural light without coloration; f.m. = moss leaves; ar. = areolae; c.b. = bacterial colonies; length of segment = 50 µm
<figure><img src="image_29.png" /><figcaption>Fig. 203: Enchytraeid faeces, detail showing a cyanobacterial colony (LN); Fig. 204: Oribatid faeces, overview (LN); Fig. 205: Oribatid faeces, detail showing melanised hyphae of *Cenococcum geophilum*, with intact walls, cut into short pieces (CP); Fig. 206: Idem, detail showing melanised hyphae (intact) and hyaline hyphae (degraded) (CP); Fig. 207: Idem, detail showing a bacterial colony embedded in dematiaceous fungal material (CP); Fig. 208: Idem, detail showing dematiaceous fungal material, Collembolan scales and bacterial colonies at the periphery (CP); Fig. 209: Idem, detail showing algal development at the periphery (CP); Fig. 210: Idem, detail showing the passage from intact melanised fungal hyphae to a coarse assemblage of strongly degraded walls (CP)</figcaption></figure>

CP = phase contrast; LN = natural light without coloration; c.a. = cyanobacterial colonies; h.h. = hyaline hyphae; h.m. = melanised hyphae; c.b. = bacterial colonies; a. = algae; length of segment = 50 µm
Oribatid faeces were much more abundant than in the overlying L₁ layer. They were isolated, deposited at the surface of decaying plant material or embedded in the basidiomycetous mycelium of *Hyphodontia* sp. Their shape was characteristic (Fig. 204). To the exception of some fragments of pine pollen or scales of Collembola, oribatid faeces were comprised of fungal material, mostly melanised hyphae of *Cenococcum geophilum*, which were cut into short fragments (Fig. 205). When hyaline and melanised hyphae were found side by side, differences in their degradation stage were striking (Fig. 206). Bacterial colonies were observed, either in micro-cavities among compacted fungal hyphae (Fig. 207) or at the periphery of oribatid faeces (Fig. 208). Unicellular algae (Chlorophyceae) were seen proliferating at the surface of some faeces. These algae were of the same type as those observed in L₁ (Fig. 71), which were responsible for the green hue of oribatid faeces, but they were far less abundant, probably due to light interception by the overlying green moss layer.

I observed in some faeces a change in the aspect of melanised hyphae of *Cenococcum geophilum*. Cell walls had lost their neat contour, being transformed in a compact amorphous mass. Transitional stages could be observed at the inside of some ageing faeces (Fig. 210) until in some faeces the totality of the faecal material became amorphous (Fig. 211). I never observed such a phenomenon in the other invertebrate groups I scrutinized. Given that no microorganism was found associated with amorphous stages of *Cenococcum* degradation, I hypothesized that this degradation resulted from non-biotic processes, maybe involving free enzymes.

Exceptionally at this level, some oribatid faeces showed that pine needles could be consumed, too (Fig. 212). The presence of needle epidermis and the absence of tracheids suggested that internal tissues were not consumed, biting by oribatid mites being not followed by tunnelling activity (but see F₁ layer).

## Springtail faeces

Faeces of Collembola were found at the surface of decaying pine needles, isolated as for oribatid faeces, but at a much lower level of abundance. Their dark colour was due to the abundance of melanised fungal material. Contrary to what has been observed in living specimens, the content of faeces found in L₂ (Figs. 213 & 214) indicated mycophagy rather than coprophagy. Hyaline hyphae were degraded while walls of melanised hyphae remained intact. Bacterial development was always intense (Fig. 215).

## Animal corpses and corpse fragments

At first it should be noted that a lot of corpses and exuviae were included in invertebrate faeces (see Fig. 189). Thus most arthropod remains were not included in the list presented in Table 6, which takes into account only cuticles and fragments of cuticles which were found free in the environment. Oribatid mites were largely dominant. Two specimens of box mites (oribatid mites with complete closure of anterior part and legs in the body) were found (Table 3), indicating that this group, although not found alive in L₂ at the time of sampling, had previously colonised it.
<figure><img src="image_30.png" /><figcaption>Fig. 211: Oribatid faeces, detail showing strongly decayed melanised hyphae of *Cenococcum geophilum* (CP); Fig. 212: Idem, detail showing fragments of pine epidermis and bacteria (LN); Fig. 213: Springtail faeces, detail showing fragmented melanised hyphae of *Cenococcum geophilum* (CP); Fig. 214: Idem, detail showing fungal material and bacteria (CP); Fig. 215: Idem, bacterial development and fungal material (CP); Fig. 216: Corpse of *Adoristes ovatus*, detail showing a network of melanised hyphae at the surface of the cuticle (CP); Fig. 217: Fly wing, detail showing melanised hyphae of *Cenococcum geophilum* (CP); Fig. 218: Corpse of *Oppia sp.*, detail showing bacterial development at the inside (CP)</figcaption></figure>

CP = phase contrast; LN = natural light without coloration; f.e. = fragments of epidemis cell; h.m. = melanised hyphae; b. = bacteria; length of segment = 50 µm
<table><caption>**Table 6.** Number of animal corpses and corpse fragments found under the binocular microscope in the L₂ layer</caption><tr><td>Fragments of arthropod legs</td><td>2</td></tr><tr><td>Unidentified mesostigmatid mites</td><td>2</td></tr><tr><td>Unidentified oribatid mites</td><td>2</td></tr><tr><td>Unidentified box mites</td><td>2</td></tr><tr><td>Adoristes ovatus</td><td>2</td></tr><tr><td>Oppia sp.</td><td>1</td></tr><tr><td>Oribatula tibialis</td><td>1</td></tr><tr><td>Platynothrus peltifer</td><td>1</td></tr><tr><td>Nanhermannia nana</td><td>1</td></tr><tr><td>Fly wing</td><td>1</td></tr></table>

As in the L₁ layer (Fig. 73) the surface of oribatid corpses was covered with a network of melanised hyphae (Fig. 216). A similar fungal development was observed at the surface of a fly wing (Fig. 217). However, chitinolysis was never registered. The inside of bodies was invaded by bacteria (Fig. 218), and protozoa could be observed, too, such as for instance the small cyst amoeba *Corythion dubium* (Fig. 219).

<figure><img src="image_31.png" /><figcaption>Fig. 219 (x 400) *Corythion dubium* (cyst amoeba) at the inside of an arthropod leg (natural light without coloration)
th. = thecamoeban cyst; length of segment = 50 µm</figcaption></figure>

Conclusions about the L₂ layer

The process of fungal decomposition of litter initiated in L₁ was pursued in L₂, mainly under the influence of *Verticicladium trifidum*, a specific decomposer of pine needles. A novel stage could be defined, characterised by a bacterial and algal (cyanobacterial) development at the inside of pine needles, as well as outside them, in the numerous foldings issued from the collapse of parenchyma. This bacterial/algal development occurred mainly to the detriment of previous fungal colonisers, which achieved the destruction of internal tissues: lignified parts of pine needles were destroyed by *Verticicladium trifidum* while pine wood was destroyed by the white rot *Hyphodontia sp*.
The successional development of bacterial communities in such a superficial layer occurred mainly through the agency of enchytraeids and several macroinvertebrates, lumbricid activity being suggested by the deposition of typical earthworm faeces. Biting and nibbling of pine needles by fauna and the tunnelling activity of enchytraeid worms caused a disruption of the hard envelope (epidermis + hypodermis) which protected pine needles from the external environment, and microbial propagules were dispersed directly at the inside of decaying needles.

What was the fate of plant organic matter once incorporated into invertebrate faeces? The present study showed that an intense microbial (bacterial, algal) development was observed at the inside of faeces. The strong compaction of the fungal material within oribatid faeces, probably stemming in local anaerobiosis, was at the origin of a particular degradation of melanised hyphae, not accompanied with bacterial development. This formation of stable amorphous organic matter (humus sensu stricto) should be better described and followed in TEM microscopy. Microbial development seemed less intense in faeces of macroinvertebrates than in those of enchytraeids. In particular the development of algae and bacteria and the stage of degradation of the plant material in macroinvertebrate faeces did not seem different from what was observed in unconsumed pine needles. This was probably due to the fact that only coarse fragments were ingested by bigger animals, which did not result in disruption of plant cell walls. Such physical limit to the impact of soil animals on the degradation of plant litter was overlooked by most authors. Enchytraeid worms, by finely comminuting plant tissues and incorporating them in loosely packed faecal material (contrary to oribatid mites which compact it), favoured further bacterial development. However, at the studied depth level, the amount of plant litter which had transited by enchytraeid guts was still very feeble.

The algal (cyanobacterial) development which had been registered at the inside of pine needles was probably favoured by the shallow water table (pseudo-gley), which maintained a high level of available moisture even in summer months, as was indicated by the presence of *Molinia caerulea*. This might also explain the abundance of enchytraeids in full summer, as well as the presence of copepods, as will be seen below. The existence of algae at the inside of closed plant organs such as pine needles should deserve a special study on their metabolism when light is absent or strongly filtered out. When examined in water, these algal colonies appeared to be perfectly transparent, without any visible pigment. I suspect them to become heterotrophs when deprived of light, and thus to play the same role as bacteria and fungi in decomposition processes.

The optical method used in the present study allowed visualizing most interactions between microbes, animals, plants and minerals taking place in the L₂ layer, although the latter component was still negligible so near the surface. Within the abovementioned limits of the method (poor recovery of macroinvertebrates, imperfect assessment of microfaunal processes, absence of replication in time and space, poor representativeness) this method, cheap and easy to perform in every laboratory equipped with a good light microscope, was able to describe poorly known processes such as those which happens at the inside of a pine needle. In addition, the choice of a restricted volume to be studied allowed an overview of the soil ecosystem at the right place and at the right scale at which most interactions between plants, microbes and animals take place, thereby escaping the terrible problem of soil heterogeneity.
THE F₁ LAYER: FRAGMENTED NEEDLES, INVERTEBRATE FAECES AND MYCORRHIZAE

<figure><img src="image_32.png" /><figcaption>**Figure 220.** The F₁ layer before fixation and transport to the laboratory</figcaption></figure>

Table 7 shows the respective volume of the gross components of the F₁ layer. Compared to the overlying L₂ layer, it should be noticed that the diversity of the observed material did not change longer.

<table><caption>**Table 7.** Volume of the different components of the F₁ layer sorted under the binocular microscope</caption><thead><tr><th>Litter components</th><th>Volume (µL)</th></tr></thead><tbody><tr><td>Pine wood</td><td>4200 ± 100</td></tr><tr><td>Moss</td><td>2700 ± 100</td></tr><tr><td>Macroinvertebrate faeces</td><td>1500 ± 20</td></tr><tr><td>Bracken fronds</td><td>150 ± 100</td></tr><tr><td>Pine bark</td><td>1200 ± 100</td></tr><tr><td>Pine twigs</td><td>1200 ± 100</td></tr><tr><td>Pine roots</td><td>800 ± 100</td></tr><tr><td>Pine euphylls</td><td>180 ± 20</td></tr><tr><td>Miscellaneous</td><td>100 ± 20</td></tr><tr><td>Male flowers of pine</td><td>55 ± 5</td></tr><tr><td>Complex assemblages</td><td>45 ± 5</td></tr><tr><td>Living fauna</td><td>4.6 – 18</td></tr><tr><td><strong>Total</strong></td><td><strong>13500 ± 100</strong></td></tr></tbody></table>
The abundant bracken litter, the presence of some strongly decayed fragments of birch leaves allowed to date approximately the fall of the plant material present in F₁ to a period extending from summer 1980 (a year before sampling date) to December 1980 or January 1981. This lapse of time embraced a peak of needle fall in September and October 1980 (Arpin et al., 1986a). The presence of euphylls and male flowers of pine (fallen in June) in F₁ was probably due to the penetration of these very small and light plant organs through loosely packed L₁ and L₂ layers, following intense rains which occurred in the course of summer 1981, previous to sampling.

The proportion of voids (73%) was only slightly less than in L2 (77%): compaction was thus still poor at 5-6 cm depth (Table 1). Tables 2, 4 & 7 showed that F₁ was the first layer where pine needles were dominant and overwhelmed moss material. However, this should not be translated in terms of litter decomposition (which would be slower for pine than for moss), because maximum needle fall occurred during the period corresponding to the deposition of F₁ material. In addition, non-linear trends in litter decomposition were evidenced by the dramatic increase of macroinvertebrate faeces, which became the third litter component in volume in F₁ (11% of the total volume of solid matter against 2% in L₂). Root development was ten times more important in F₁ compared to L₂ (6% in place of 0.6%). The biovolume of living fauna, within the abovementioned limits of its calculation, was also higher than in L₂.

Pine needles were fragmented under the action of macrofauna (slugs, earthworms, woodlice, which were identified by their faeces) and mesofauna (enchytraeids, oribatid mites, which were directly observed). Oribatid mites deposited their typical egg-shaped faeces at the inside of needles (Fig. 221), external tissues remaining intact (their faeces could be observed only by breaking needles with scissors). Enchytraeids penetrated also pine needles but most of time only the anterior part of the body plunged into them (Fig. 222) and as a consequence they defecated mainly in the outside. Some sciarid larvae were also observed penetrating pine needles, in the same manner as enchytraeids. Thus, several widely different animal groups shared the same decaying plant material as food resource and cohabited in a small volume of litter, without any segregation in space or time. Such observations were in disagreement with commonly held views on competitive exclusion and niche separation (Barbault, 1981) but they were supported by the direct observation of multi-species cultures (Christiansen, 1967). When not ingested by fauna fragmented needles were embedded in the same woolly mycelium described in the L₂ layer. Observations on pine ectomycorrhizae showed that this mycelium, which was comprised of the ascomycete *Cenococcum geophilum* and the basidiomycete *Hyphodontia sp.*, as mentioned above, were linked to the two ectomycorrhizal types which were dominant in F₁. Most often, external signs of previous fungal colonisation (pycnidia, ascocarps, diaphragms, conidiophores) had disappeared from the surface of pine needles, due to biting and nibbling by fauna. However, diaphragms of *Lophodermium pinastri* were still sometimes observed, together with conidiophore remains of *Verticicladium trifidum*. Unidentified black sclerotia which had been observed at the surface of some pine needles in L₂ were also observed in F₁, but rhizomorphs of the basidiomycete *Marasmius androsaceus* were absent. It should be also noticed that the past activity of the latter fungus was not registered on sectioned pine needles, pointing to a strongly seasonal development of this fungus (winter to spring, as mentioned above), which was probably not able at that time to colonize needles at a too advanced stage of decomposition. Decaying needles were strongly intermingled with other components of the F₁ layer. Pine ectomycorrhizae were often seen tightly appressed against fragments of pine needles, leafy shoots of moss and animal faeces, these assemblages being embedded in turn in fungus mycelium. Growing roots seemed to preferentially penetrate organic assemblages of decaying plant material, making their isolation particularly tedious.
<figure><img src="image_33.png" /><figcaption>Fig. 221: Oribatid faeces at the inside of a pine needle; Fig. 222: Two enchytraeid worms (arrows) penetrating a pine needle, with deposited their faeces; Fig. 223: Type A excrement (slug?); Fig. 224: Type B excrement (woodlouse?); Fig. 225: Type C excrement (earthworm); Fig. 226: Long fine root of pine colonised by *Cenococcum geophilum* in the 'root hair' zone and with two ectomycorrhizae of the same fungus; Fig. 227: Long fine root of pine with growing apex and two different types of ectomycorrhizae (black ectomycorrhiza of *Cenococcum geophilum* and orange-brown ectomycorrhiza of *Hyphodontia sp.*); Fig. 228: Pink basidiomycetous ectomycorrhiza</figcaption></figure>

length of segment = 1 mm
At last, all pine needles appeared to be strongly collapsed, with sometimes only the external envelope (hypodermis + epidermis + cuticle) remaining.

Moss leafy shoots seemed to be more microbially attacked than in the above $L_2$ layer. They were firmly appressed to each other, sometimes mixed with bracken leaflets, and abundantly covered (or intermingled) with animal faeces. As well as other plant material they were embedded in the at once ubiquitous fungal mycelium. Moss stems exhibited often a dark red colour, which had not been observed in $L_2$, and some of them were empty. In the latter case they were translucent and light coloured. White rhizomorphs were seen protruding from some decaying moss stems. Ramified moss stems were often seen totally deprived of leaves, probably following peeling by fauna.

Three types of macroinvertebrate faeces were observed in $F_1$. The A type (Fig. 223) was represented by more or less folded centimetric folded strands which were made of long fragments of pine needles which had been seemingly swallowed entire. The ingested plant material did not seem to have been transformed during gut transit. By their size and shape I interpreted these excrements as slug faeces, even though these animals were present in the study site but not in the studied sample. The B type (Fig. 224) was represented by truncated, flattened millimetric cylinders. Under the dissecting microscope, they appeared to be comprised of very small, compressed fragments of pine needles. Their colour was more or less dark brown, but never blackish. Fungal hyphae were observed under the light microscope, but they could come from secondary colonisation. By their size and shape these faeces were attributed to woodlice, despite of the fact that the two *Trichoniscus* larvae which were found in the sample did not exhibit a similar composition in their gut contents. The C type (Fig. 225) was by far the most abundant type of macroinvertebrate faeces found in the $F_1$ layer. Their colour was blackish and they were strongly convoluted without any definite external shape. Their content was finely ground, contrary to types A and B. Type C faeces were often found appressed against plant material, probably because of their pasty nature when excreted and of the movements of animals which pushed their way through litter. They were attributed to epigeic earthworms, in particular to *Dendrobaena octaedra* (Savigny) which was dominant in the site (Bouché, unpublished data). Other earthworm faeces were found in the sample, but they were not accounted for, given their poor representativeness. They were mineral, friable, white or light gray coloured. These excrements were produced either by another earthworm species or by *Dendrobaena octaedra* when dipping in underneath horizons: some authors like Martinucci & Sala (1979) classified this species as a 'weak burrower'.

Faeces of mesofauna (microarthropods, enchytraeids) were either dispersed at the surface of decaying material (oribatids, springtails) or agglomerated at the inside of needles or bark pieces (box mites, sometimes enchytraeids) or outside of them (enchytraeids, sciarid fly larvae). Despite of the intense activity of these animals (in particular enchytraeids and box mites) the total volume of faeces they produced was largely less than that of macroinvertebrates, in particular type C (earthworms).

The root network in $F_1$ was only comprised of fine roots. Long roots exhibited a growing apex, with an elongation zone (sometimes with absorbing hairs below it). They bore short ectomycorrhizal roots (Fig. 226). While the ectomycorrhizae present in $L_2$ were always of the coralloid orange-brown type, a wider variety of ectomycorrhizal types was depicted in $F_1$, probably because fine roots were older in $F_1$ compared to $L_2$ (Figs. 226 to 228). The dominant type was the black ectomycorrhiza of *Cenococcum geophilum* [= *Cenococcum graniforme* (Sow.) Ferd. & Winge]. This ascomycete is only known as a sterile mycelium (sometimes called 'jet-black mycelium') which exhibits many analogies with an *Elaphomyces* according to Trappe (1971). It produces spherical
sclerotia of varying size, which were found in abundance in the studied sample (Fig. 229). Long straight black hyphae protruded from ectomycorrhizae (Figs. 226 & 227) and sclerotia (Fig. 229) of *Cenococcum geophilum*, permeating the decaying plant material. Earthworm faeces (type C) were also a favourable seat for the development of this fungus (Fig. 230). Ectomycorrhizae of *Cenococcum geophilum* were short, dichotomous (Fig. 226) or monopodial (Fig. 227), and the fungus might also colonize the long root, in the zone normally carrying root hairs (Figs. 226 & 227). Cohabitation with the coralloid orange-brown type was observed, either on the same long root but on distinct short roots (Fig. 227) or even on the same short root. Most often, the growth of *Cenococcum geophilum* was slower than that of long roots, letting uninfected root apices but, as has been already shown by Wilcox (1968), it happened infrequently that the apex was colonised by the fungus, and root growth was checked (Fig. 231). My observations did not allow discerning which event (growth arrest of the root or fungus infection) occurred first. The coralloid type was always associated with a white woolly mycelium (Fig. 227) of the same type as the one embedding decaying moss shoots, pine needles, etc... The third ectomycorrhizal type, much more infrequent, was represented by pink, thick, velvety dichotomous ectomycorrhizae (Fig. 228), without any mycelial association visible under the dissecting microscope.

## Decaying pine needles

### Fungal mantle around pine needles

Most pine needles were embedded in a fungal mantle which was comprised of four distinct types, three of which proved to be associated with pine ectomycorrhizae. The most abundant type was the mycorrhizal ascomycete *Cenococcum geophilum*, in its typical aerial form (Fig. 232). This fungus was present in $L_1$ and $L_2$, but its mycorrhizal nature was verified in $F_1$, where it was associated with black ectomycorrhizae (Figs. 226, 227 & 231) and sclerotia (Fig. 229). The mycelium of *Cenococcum geophilum* was accompanied by the secondary mycelium of the white-rot basidiomycete *Hyphodontia sp.*, hyphae of which were encrusted with oxalate crystals (Fig. 233) or not (Fig. 232). Observations done in $F_1$ (see below) showed that this fungus, although definitely a white rot as already shown in $L_2$, (Figs. 120 & 121), was associated with the coralloid orange-brown ectomycorrhizae of pine. The third type was intermingled with the secondary mycelium of *Hyphodontia sp*. It was made of smooth hyaline hyphae of 10 µm diameter, at the inside of which intra-hyphal regrowth (hyphae of 3 µm diameter) was frequently observed (Fig. 233). Despite of the fact that clamp connections were not found on its hyphae in the $F_1$ layer, I thought that the same fungus had been observed embedding moss leaves in $L_2$, similarly in the company of *Hyphodontia sp*. (Fig. 131). This unidentified fungus was not found in association with pine ectomycorrhizae, but it cannot be excluded that such big smooth hyphae could differentiate from ordinary hyphae in the mycelium of *Hyphodontia sp*. Two arguments were in favour of this hypothesis, (i) hyphae issued from intra-hyphal regrowth of the third type were morphologically similar to hyphae of *Hyphodontia sp*. (when not encrusted with oxalate crystals), and (ii) both fungal types were always found tightly intermingled. The fourth type was rarer. It was made of smooth hyphae, somewhat larger (4 µm) than those of *Hyphodontia sp*. The same mycelium was found in association with pink velvety ectomycorrhizae of pine (Fig. 228).
<figure><img src="image_34.png" /><figcaption>Fig. 229: Sclerotium of *Cenococcum geophilum*; Fig. 230: Type C (earthworm) excrement from which *Cenococcum geophilum* hyphae were protruding; Fig. 231: Long fine root of pine the growth of which was arrested and the apex colonised by *Cenococcum geophilum*; Fig. 232: Pine needle, section showing from right to left fine hyphae of *Hyphodontia sp.*, coarse hyphae of *Cenococcum geophilum*, bacterial development between hyphae, epidermis and hypodermis transformed in a melanised fungal stroma (CP); Fig. 233: Idem, showing fine hyphae of *Hyphodontia sp.* encrusted with oxalate crystals and smooth coarse hyphae with intra-hyphal regrowth (CP); Fig. 234: Idem, showing bacteria in way to be degraded (CP); Fig. 235: Idem, showing transfusion tissue transformed into a pulp, with bacterial development and brown spherules (CP); Fig. 236: Idem, showing a layer of unicellular algae just under the hypodermis, on both sides of a stomata (CP)</figcaption></figure>

CP = phase contrast; length of segment = 1 mm (Figs. 229-231) or 50 µm (Figs. 232-236)
A bacterial development was observed in the fungal mantle embedding decaying pine needles (Fig. 232). Sometimes hyphae of *Cenococcum geophilum* were seen penetrating decaying fungal material, maybe corresponding to a successional process within the fungal mantle. The presence of another fungus, growing as a parasite on this fungus mantle, should be noted, too. It fructified in the form of tiny white funnels, the wall of which was made of firmly appressed hyphae, replaced at the inside by short free hyphae bearing conidia. Unfortunately they could not be sectioned correctly in order to study conidiogenesis. It could be possibly a *Myrothecium* sensu lato (Morelet, personal communication).

*Inside of pine needles*

The constant presence of fauna at the inside of pine needles found in F₁ must be highlighted, even when needles did not exhibit any sign of nibbling or penetration by invertebrates. Microfauna were always observed in sectioned needles, in the form of protozoa (naked amoebae, flagellates), rotifers or nematodes. Cyst amoebae, although frequently observed at the surface of bracken leaflets, moss leaves and pine needles, were never observed at the inside of pine needles except when these had been previously tunnelled by mesofauna. The decay stage which pine needles reached in F₁ was thus characterised by the ubiquitous action of a variegated fauna. Less generally, mycorrhizal fungi were seen penetrating pine needles, more especially *Cenococcum geophilum*. Non-mycorrhizal basidiomycetes were also observed at the inside of pine needles, but only in the form of dead hyphae, testifying for previous fungal colonisation. Most often, and this was true for *Cenococcum geophilum*, too, fungal hyphae were dead or senescent (no staining by methyl blue, no opacity under phase contrast), while bacteria or more exceptionally micro-algae (Chlorophyceae) were found alive. A replacement of fungi by bacteria had been already observed in L₂, but now the degradation of bacteria was observed, too (Fig. 234), which might indicate a check of bacterial colonisation in F₁. The same for chroococcal algae (cyanobacteria), which were abundant at the inside of pine needles in L₂, and in way to disappear in F₁.

* Needles attacked by *Verticicladium trifidum*, but not by *Cenococcum geophilum*

The presence of *Verticicladium trifidum* was attested by its hyaline, thick-walled hyphae, which were observed in a senescent state in xylem tracheids, or by its typical stromata developed at the inside of host tissues. Some needle fragments were observed, at the inside of which mesophyll and endodermis had been entirely transformed into melanised fungal tissues. These were devoid of cytoplasm at the time of sampling but their cell walls were still intact. Epidermis and hypodermis could be also transformed into a melanised fungal stroma (Fig. 232). In all cases of previous attested colonisation by *Verticicladium trifidum*, the transfusion tissue was transformed into a pulp, where the only more or less recognizable plant elements were bordered pit areolae, with fuzzy contours (Fig. 235). Bacterial development was intense (Fig. 235) except in the unique case observed where the fungus was still active (although senescent). Some traces of other types of fungal colonisation could subsist, too, such as empty hyphae and diaphragms of *Lophodermium pinastri*. Needles where the
development of *Verticicladium trifidum* was not arrested by the faunal transformation of pine needles were characterised by further cellulolysis in the central cylinder (to the exception of xylem) and the development of fungal stromata which replaced progressively epidermis, hypodermis, mesophyll and endodermis. Even when internal tissues were further invaded by bacteria, it should be noticed that the degradation of host cell walls was not caused by bacteria but rather by *Verticicladium trifidum*, since similar figures were observed in L₂ in the absence of bacteria (Fig. 76).

Other phenomena should be noticed, too, the importance of which and the assignment to *Verticicladium trifidum* activity were more disputable since they not repeatedly observed: the presence of big-sized brown spherules in the transfusion tissue of a pine needle (Fig. 235), and the development of a unicellular algal (Chlorophyceae) layer just under the hypodermis, which had been transformed into a melanised stroma (Fig. 236). The latter pattern, observed on a wide area of a unique needle fragment, had not been registered in the overlying L₂ layer.

* Needles attacked by *Cenococcum geophilum*

This ascomycetous ectomycorrhizal fungus was observed embedding needle fragments in F₁, where it was accompanied by basidiomycetous ectomycorrhizal fungi. However, it was also often seen penetrating pine needles, where the morphology of its mycelium was quite different from that of the aerial form. Contrary to *Verticicladium trifidum*, which invaded in a progressive manner the totality of internal tissues of pine needles, *Cenococcum geophilum* formed a dense network of entangled hyphae in the part of the mesophyll which was located just under the hypodermis (Fig. 237). Given that remains of pine tissues were not included within the mycelial network of *Cenococcum geophilum* but were pushed aside, it was thought highly probable that this fungus did not contribute to the degradation of pine needles. Entangled hyphae were convoluted, of smaller section (2 µm) than the aerial form (4 µm) and were less impregnated with melanin (grey vs jet black for the aerial form). Chlamydospores (Ø 4-7 µm) were observed from place to place. Although strongly different from needle-embedding hyphae, entangled hyphae found at the inside of decaying pine needles were attributed unambiguously to *Cenococcum geophilum*. Some zones were found, where both forms cohabited in the same mycelium and were connected to each other (Fig. 238). Both types of hyphae fall within the wide morphological variability of *Cenococcum geophilum*, which has been described in agar and liquid culture by Hatch (1934) and Mikola (1948), respectively. The morphological variety observed within the same fungal strain (see also strain varieties described by Mikola) is also at the origin of the numerous names given to this fungus in ancient studies (Hatch, 1934). The entangled hyphae of *Cenococcum geophilum* observed between the mesophyll and the hypodermis appeared always devoid of any cytoplasm, some were even in way to be degraded (Fig. 239). However, no bacterial development was registered. The mesophyll and the central cylinder showed signs of previous fungal colonisation, but typical melanised hyphae of *Cenococcum geophilum* were observed in some xylem tracheids. In this case, I suspected that needles were first fragmented by fauna then the sectioned central cylinder was colonised directly at the level of a break, since no hyphal connection was found with the mycelium which developed under the hypodermis. Needles colonised by *Cenococcum geophilum* seemed highly palatable to box mites, since tunnelling by these animals always occurred in needles previously colonised by the ascomycete. However, I observed that box mites ingested only internal tissues of decaying pine needles, not the melanised dense hyphal network
<figure><img src="image_35.png" /><figcaption>Fig. 237: Pine needle attacked by *Cenococcum geophilum*, section showing a network of entangled hyphae in the portion of the mesophyll located just under the hypodermis (CP); Fig. 238: Idem, showing a mixed network of large straight and small convoluted hyphae (CP); Fig. 239: Idem, showing entangled hyphae in way to be degraded (CP); Fig. 240: Coralloid orange-brown pine ectomycorrhiza with *Hyphodontia sp.*, detail of a hypha of from the extramatrical mycelium showing the passage from smooth wall to wall encrusted with oxalate crystals (CP); Fig. 241: Idem, showing a cystidium born by a hypha (CP); Fig. 242: Black pine ectomycorrhiza with *Cenococcum geophilum*, detail of the extramatrical mycelium showing melanised hyphae with encrusted walls (LN); Fig. 243: Idem, detail showing the passage from hyaline to melanised hypha on the same mycelium (CP); Fig. 244: Pink pine ectomycorrhiza, detail of the prosenchymatous fungal mantle (CP)</figcaption></figure>

CP = phase contrast; LN = natural light without coloration; length of segment = 50 µm
produced by *Cenococcum geophilum*. The intense faunal activity which took place within needles colonised by *Cenococcum geophilum* made them rapidly emptied, and further colonised by other mycorrhizal fungi. However, mycorrhizal basidiomycetes were never observed at the inside of needle tracheids, they just colonised galleries created by tunnelling animals.

Brown spherules similar to what has been observed in other decaying needles in F₁ (Fig. 235) and L₂ (Fig. 104) were found only once. Their interpretation as polyphenol-protein complexes precipitating at the death of cells, a common phenomenon described by Handley (1954), is rather problematic, since they were observed only on a few needles. Thereby the question of their origin remains open.

The most prominent character of needles colonised by *Cenococcum geophilum* was the incompatibility of this fungus with bacterial and algal development. It must be highlighted that the degradation of bacterial cells (Fig. 234) was observed in a needle colonised by *Cenococcum geophilum*, a mycorrhizal ascomycete known for its production of antibiotics (Grand & Ward, 1969). However, it did not seem to be involved in the degradation of pine tissues, cells of which (in particular tracheids) were penetrated without any alteration of their wall. *Cenococcum geophilum* seemed also incompatible with *Verticicladium trifidum*, since needle fragments colonised by the former did not seem to have been previously colonised by the latter. The fact that hyphal walls of *Cenococcum geophilum* seemed devoid of cytoplasm might suggest that colonisation by this fungus took place during previous autumn and/or winter, while that by *Verticicladium trifidum* was more recent: viable hyphae of this fungus were visible in F₁ and were very abundant, with concomitant fructification, in L₂ and L₁ (seemingly deposited in spring and summer). Curiously, data on the seasonal development of *Cenococcum geophilum* mycelium were lacking in literature, only its mycorrhizae being studied in detail.

## Pine roots and mycorrhizae, mycorrhizal fungi

Many authors described anatomical characters of pine ectomycorrhizae, either in light (Hatch & Doak, 1933; Hatch, 1934; McDougal & Dufrenoy, 1944; Robertson, 1954; Fassi & De Vecchi, 1962; Marks, 1965; Fassi & Fontana, 1966; Scannerini & Palenzona, 1967; Wilcox, 1968; Fortin et al., 1980) or electron microscopy (Foster & Marks, 1966; Marks & Foster, 1967; Scannerini, 1968; Hofsten, 1969). Extramatrical mycelia, although neglected by most authors, were described, in particular in the rhizosphere (Foster & Marks, 1967) and when formed of rhizomorphs (Duddridge et al., 1980; Foster, 1981; Brownlee et al., 1983). This is the reason why only poorly described features of mycorrhizae as well as root decomposition patterns will be illustrated here.

### Living roots

* Coralloid orange-brown ectomycorrhizae
These mycorrhizae, already observed in L₂, were characterised by repeated dichotomous growth, stemming (not always, but often) in coralloid masses. The observation of the fungal mantle in tangential section allowed me to discern its prosenchymatous nature: it was formed of cemented hyphae, not separable with forceps. The thickness of the mantle was highly variable, sometimes of only one cell layer. It disappeared towards the apex of mycorrhizal branches. Contrary to external colonisation, internal colonisation was pronounced, even towards apices, with a 1-2 cell thick fungal Hartig net separating hypertrophied cortical cells of pine. Starch grains abounded at the inside of hypertrophied cortical cells. The peripheral cortical zone, just under the fungal mantle, was comprised of two layers of 'tannin' cells. In the peripheral zone of the fungal mantle the prosenchyma passed progressively (hyphae were more and more discernible, not cemented) to a woolly extramatrical mycelium made of smooth hyphae (3 µm diameter) with clamp connections, i.e. to the dicaryotic secondary mycelium of a typical basidiomycete. Within the limits of this purely morphological classification, coralloid orange-brown ectomycorrhizae can be classified in the type Aₑ of Dominik (1969).

The link between the smooth extramatrical mycelium of coralloid orange-brown ectomycorrhizae, the encrusted hyphae embedding pine needles and moss shoots and the white-rot fungus identified as *Hyphodontia sp.* (arguta-pallidula-alutaria group) was indirectly established through the observation of several anatomical characters. First, hyphal diameter, size and shape of clamp connections, ramification angle and wall thickness were identical between both mycelial forms. Second, the passage from smooth to encrusted zones was observed along the same hyphae (Fig. 240), pointing to versatility of oxalic acid excretion across hyphal walls. Third, a lagenocystidium was observed in the extramatrical mycelium of coralloid orange-brown ectomycorrhizae (Fig. 241), quite identical to cystidia observed in decaying branchwood (Figs. 120 & 121). This fungus was thus suspected to be both mycorrhizal and saproxylic, a behaviour which is not uncommon in Corticiaceae (Käärik & Rennerfelt, 1957; Vozzo & Hacskaylo, 1971).

* Black ectomycorrhizae

These jet-black ectomycorrhizae, unambiguously produced by the sterile ascomycete *Cenococcum geophilum*, corresponded to the Gᵃ type of Dominik (1969). They were described under various names by many authors such as Hatch (1934) under *Mycelium radicis nigro-strigosum*, Mikola (1948), Trappe (1971), Kiffer (1974) and Voiry (1981) under the name *Cenococcum graniforme*. Marks & Foster (1967) described quite identical black mycorrhizae but without naming the fungus.

The external morphology of black ectomycorrhizae was variable. They were either club-like (Fig. 227) or slightly dichotomous (Fig. 226) but never ramified. Straight hyphae merging at right angle from the mycorrhiza were typical of the 'jet-black mycelium' of Hatch (1934). In light microscopy, the thick fungal mantle appeared pseudo-parenchymatous, being formed of four layers of septate melanised hyphae, cemented in palisade arrangement. Contrary to the previous coralloid type, the thickness of the fungal mantle increased towards the apex. Melanisation of mantle cells decreased towards the interior of the mycorrhiza. The hyaline Hartig net was thinner than in the coralloid type, being formed of only one cell layer. Only one layer of host 'tannin' cells was formed under the mantle. Below it there was an external zone formed of dead parenchymatous cells, followed at the inside by alternating living and dead cortical cells. Thereby internal colonisation by the fungus was less
pronounced than in the coralloid type, but the host seemed more affected by *Cenococcum geophilum*
than by *Hyphodontia sp.*

I often observed, in the proximal part of black ectomycorrhizae where the fungal mantle of
*Cenococcum geophilum* was absent, an imperfect fungal mantle formed by the basidiomycete
*Hyphodontia sp.*, hyphae of which were not cemented and bore clamp connections. When sectioned at
this level (identified by a constriction of the cortex), black mycorrhizae showed the 2-cell Hartig net
and the double layer of ‘tannin’ cells typical of orange-brown coralloid mycorrhizae, testifying for
internal colonisation of the root by *Hyphodontia sp*. The coexistence of two fungi on the same
mycorrhizal root has been already described by Marks & Foster (1967), who explained it by a
successional process. The coexistence of *Cenococcum geophilum* and *Hyphodontia sp.* on the same
short root was probably made possible by their different ratio of external and internal colonisation.

The mycelium of *Cenococcum geophilum* was comprised of melanised hyphae with thick walls (generally diameter 4-5 µm) most often smooth (Figs. 232 & 238) but sometimes encrusted with minute bumps (Fig. 242). Although not the rule (contrary to the inside of pine needles), the abrupt passage to melanin-free and thinner hyphae has been observed in the aerial mycelium, only intra-hyphal walls remaining impregnated with melanin (Fig. 243).

* Pink ectomycorrhizae

These mycorrhizae, of velvet aspect and always dichotomous (Fig. 228), bore a thick mantle
which was strongly variable in its anatomical features, making their classification difficult according
to Dominik (1969). When the mantle was pseudo-parenchymatous (true plectenchyma) they belonged
to the F<sub>e</sub> type but when the mantle was prosenchymatous, with recognizable hyphae (Fig. 244), they
belonged to the B<sub>e</sub> type. This pinpoints the awkward generalization of Dominik’s classification.
Melanised hyphae of *Cenococcum geophilum* were sometimes observed penetrating the mantle of pink
ectomycorrhizae. The Hartig net was strongly developed, but only one layer of ‘tannin’ cells was
observed in the cortex. Just below the ‘tannin’ layer brown spherules were often observed at the inside
of cortical cells (Fig. 245), quite similar to those observed at the inside of pine needles (Figs. 104, 105
& 235). The mantle became thicker towards the apex, but Hartig net and ‘tannin’ cells disappeared,
pointing to external colonisation only in the apical part of pink mycorrhizae.

Tangential sections revealed the prosenchymatous nature of the mantle, with numerous clamp
connections (Fig. 246). Pink mycorrhizae were smooth, thus not in direct association with an
extramatrical mycelium. However some aerial hyphae exhibited the same anatomical characters as
those hyphae found in the mantle: presence of clamp connections, smooth thin hyphal walls of
diameter 4 µm, i.e. more than those of *Hyphodontia sp*. The same mycelium was found embedding
dead moss in the L<sub>2</sub> layer (Fig. 131), and decaying plant debris and animal faeces in the F<sub>1</sub> layer (Fig.
255), in association with thin encrusted hyaline hyphae of *Hyphodontia sp*. and thick melanised
hyphae of *Cenococcum geophilum*.

* Long roots
<figure><img src="image_36.png" /><figcaption>Fig. 245: Pink pine ectomycorrhiza, detail of the cortex, showing brown spherules in cortical cells (CP); Fig. 246: Idem, tangential section through the prosenchymatous fungal mantle with clamp connections (arrows) (CP); Fig. 247: Living long root of pine, section through the cortex showing a Hartig net (CP); Fig. 248: Decaying long root of pine, longitudinal section through the central cylinder showing the intracellular development of basidiomycetous hyphae (with clamp connections) of *Hyphodontia* sp. (CP); Fig. 249: Sclerotium of *Cenococcum geophilum*, overview of the section (CP); Fig. 250: Idem, detail if the wall (CP); Fig. 251: Idem, detail of the heart (CP); Fig. 252: Decaying pine twig, longitudinal section through the wood showing a soft-rot type of fungal attack of tracheids (BMSL)</figcaption></figure>

CP = phase contrast; BMSL = methyl blue-Sudan III-lactophenol coloration; length of segment = 50 µm
Hair roots were rarely observed: they were located just behind the growth (elongation) zone. In this case no sign of fungal colonisation was visible, in particular there was neither Hartig net nor 'tannin' cells.

Even in the absence of externally visible sign, most long roots were colonised by mycorrhizal fungi. The internal development of *Hyphodontia sp.* was not accompanied by the formation of a mantle (except when a very thin fungal mantle was formed) but the Hartig net was present, like in true ectomycorrhizae (Fig. 247). The longitudinal penetration of this fungus was observed in cortical cells and in the endodermis, but the central cylinder was intact, with nuclei well visible within parenchymatous cells. No 'tannin' cell layer was observed in the cortex of long roots colonised by *Hyphodontia sp.*

The colonisation of long roots by *Cenococcum geophilum* was attested by the presence of a black mantle, with its typical palisade assemblage of fungal hyphae (like in black mycorrhizae), or by the presence of free hyphae running longitudinally along the root, giving it a brown colour (which could be erroneously interpreted as a sign of decay). At the inside of the root, the Hartig net was well developed, but cortical cells were not hypertrophied and 'tannin' cells were absent. A root, the apex of which had been colonised by *Cenococcum geophilum* (Fig. 231), showed under the fungal mantle an intact meristem, with numerous nuclei and the total absence of any internal colonisation by the fungus. I interpreted this pattern as the result of an arrest of growth of the pine root, maybe temporary, which was followed by superficial overgrowth of its apex by the mycorrhizal fungus. Check of the growth of the root was attested by the lack of the typical conical shape of the apex which was observed when roots were actively growing (Fig. 227).

It must be highlighted that mycorrhizal colonisation did not affect only true mycorrhizae (more or less transformed short roots) but also the long roots which bear them (Marks & Foster, 1973). In the case of *Hyphodontia sp.*, intracellular hyphae were observed in the cortex, which could be hypothesized to affect the uptake of water and nutrients by long roots. Robertson (1954) showed that the internal colonisation of short roots by mycorrhizal fungi did not spread from the mantle but rather from previously infected long roots. The colonisation of long roots by ectomycorrhizal fungi (in our case by *Hyphodontia sp.*) strongly resembles what has been described under the term 'ectendomycorrhizae' by Wilcox (1968, 1971).

## Decaying roots

* Short roots

Decay patterns were observed only on coralloid orange-brown ectomycorrhizae. They turned neat brown. In all stages of decomposition I observed, no bacterial development was visible in internal (host) tissues. In particular, the cortex remained intact with its typical Hartig net and 'tannin' cells. Only the fungal mantle seemed affected by decay. Several stages of decomposition of the fungal mantle were observed, (i) mantle hyphae were devoid of cytoplasm, the contour of external hyphae
becoming fuzzy, indicating decay of their walls, (ii) the fungal mantle was the seat of an intense
bacterial and algal development, hyphae becoming hardly recognizable, (iii) concomitant or
succeeding to it, a stage where the mantle was invaded by other fungi, among which the
basidiomycetous mycelium associated with pink ectomycorrhizae.

* Long roots

The incipient decay of long roots was characterised by the development of the secondary mycelium of *Hyphodontia sp.* in the central cylinder (Fig. 248). Some decaying long roots showed a joint development of *Hyphodontia sp.* and *Cenococcum geophilum*. The latter fungus was also observed at the surface of the decaying root, with its hyphae embedding it. The first stage of long root decomposition seemed to be the development in the stele of ‘mycorrhizal’ fungi, the only stage observed in this layer.

It must be noticed that, to the light of present observations, long roots and short roots seemed
to behave quite differently concerning mycorrhization and processes associated with death of the root.

* Sclerotia of *Cenococcum geophilum*

The presence of sclerotia of *Cenococcum geophilum* has been widely attested in dysmoder
humus, whether called ‘mor’ or ‘raw humus’ by authors (Ferdinandsen & Winge, 1925; Meyer, 1964).
In the present sample, they were in the form of near perfect spheres, of varying size, from which many
hyphae radiated (Fig. 229). When sectioned they showed a three-layered wall and a more or less
homogeneous heart (Fig. 249). The wall (Fig. 250) was comprised of (i) a 1-cell external layer, with
thin melanised cell walls [analogous in tangential view to the mycorrhizal mantle according to Trappe
(1971)], (ii) a 7-10-cell medium layer, arranged in circular files of cells, the walls of which were thin
but less melanised than those of the external layer, and (iii) an internal 10-cell layer made of thick-
walled, deep black cells with strongly reduced lumina and poorly observable cell limits. The internal
layer was responsible for darkness and toughness of sclerotia. The sclerotial heart (Fig. 251) was made
of radial files of cells, with thin slightly melanised walls. Meatuses between heart cells were filled
with melanised wall material. I did not find hollow sclerotia of *Cenococcum geophilum*, but
Ferdinandsen & Winge (1925) noted that this phenomenon was strongly seasonal, preceding the
‘germination’ of sclerotia in spring.

Pine twigs

Preliminary remarks