Bulk isolation of basidiospores from wild mushrooms by electrostatic attraction with low risk of microbial contaminations

Mushroom collection and identification

Wild mushrooms were collected as found on the North Campus of the University of Göttingen and neighbouring areas of the village of Göttingen–Weende from 09.2011 to 09.2014. Mushrooms were photographed prior to harvest using a Cannon IXUS 115 H5 digital camera (12.1 megapixels; Canon, Krefeld, Germany). Harvested mushrooms were transported into the lab and photographed again, using a ruler as size marker to allow cap size estimations. Ecological parameters of mushrooms’ growth (biotope, substrate, host trees) and morphological characters of the mushrooms (of stipes, caps, veils, lamellate hymenia or pores and, crucially in the identification process, of spores) were recorded for species determination. Basidiospores (sizes, colour and shapes) were observed under an Axioplan 2 imaging microscope (Carl Zeiss, Göttingen, Germany), photographed by a computer-linked Soft Imaging ColorView II Mega Pixel digital camera and analysed in size with the AnalySIS^® software program (Soft Imaging System, Münster, Germany). Averages of spore sizes of collected specimens were determined from usually 5–20 spores. The field guides of Breitenbach and Kränzlin (1986, 1991, 1995), Bresinsky and Besl (1990), Flück (1995), Dähncke (2001) and Gerhardt (2010) and the Coprinus pages by Uljé (http://www.grzyby.pl/coprinus-site-Kees-Uljee/species/Coprinus.htm) were used in species identifications and the MycoBank database (http://www.mycobank.org/) was considered for current species names and higher classification.

Coprinopsis cinerea mushrooms were all of strain AmutBmut (FGSC25122) and produced in the lab on artificial YMG/T medium (4 g yeast extract, 10 g malt extract, 100 mg tryptophan, 10 g agar) under standard fruiting conditions (Granado et al. 1997). A coincidental mushroom of Leucocoprinus birnbaumii was collected from a flowerpot in a student office of the institute.

Basidiospore harvests

Any dirt and noticed animals were removed from collected mushrooms. Caps were carefully separated from stipes with a sterile razor blade and forceps. Depending on their diameter (abbreviated by Ø throughout this work) or on the cap height of mushrooms which never fully open their umbrellas (i.e. Coprinopsis picacea, Coprinus comatus), caps were kept intact or sliced into 2, 4 or more equally sized portions. In cases of mushrooms with thick fleshy pilei (Amanita strobiliformis, Coprinopsis atramentaria, all Boletales but Hygrophoropsis aurantiaca) the upper gill-less or tube-free pileus parts of caps were sliced off in order to generate sufficient free air space above mushrooms during the experiments in the Petri-dishes. Caps up to ca 4–5 cm in diameter or parts of caps in case of larger mushrooms (Ø > 4–5 cm) were laid upside-down onto sterile wet paper tissue in individual sterile plastic Petri-dishes (polystyrene, 9 cm Ø, with cams, REF 82.1473; Sarstedt, Nümbrecht, Germany), sterile standard glass Petri-dishes (9 cm Ø) or higher sterile glass dishes (9 cm in Ø, 3.2 cm in height; used for Armillaria solidipes, Pholiota squarrosa and H. aurantiaca mushrooms) covered by a lid of a plastic Petri-dish. Care was taken to ensure that there was at least 0.5 mm free airspace in Petri-dishes above the mushrooms, while the airspace between mushrooms and plastic lids on higher glass dishes were between 1 and 2 cm. Dishes were stored for a few hours to overnight (up to 18–20 h) on a bench at room temperature (RT) for spore ejection. Patterns of spores adhering to the plastic lids were photographed using a Stemi 2000-C binocular (Carl Zeiss, Göttingen, Germany) connected to the Soft Imaging ColorView II Mega Pixel digital camera. Spores attached to the lids were washed off with 200 µl sterile water or with 200 µl of sterile 0.1% Tween 80 and counted using a hematocytometer.

Directional effects of mushrooms on spore release were observed in experiments of two distinct set-ups. Fruiting bodies of a same size and age or defined parts of fruiting bodies of a species were incubated in parallel in Petri-dishes on wet tissue paper in either natural direction or in upside-down position (Experimental Set-up 1). In other experiments in order to avoid a direct contact with the wet paper tissue, mushroom samples were attached to the bases of plastic Petri-dishes by sticking them with their cap surface into a layer of sterile hand-hot water agar (1%) so that the cap surfaces touched the bottoms of the respective Petri-dishes. Petri-dishes were then incubated either in up-right position or upside-down (Experimental Set-up 2). Spores were harvested after 18 h incubation at RT either from the wet paper tissues (Set-up 1) or from the surface of the plastic lids of Petri-dishes (Set-up 2).

Two different strategies were also followed up to observe spore release over the time. First, equally sized and aged mushrooms or parts of mushrooms of a species were in parallel incubated upside-down on wet tissues in Petri-dishes at RT. At defined time points, spores were harvested from selected individual samples and counted. Data from the different individual samples for the different time points were compared (Experimental Set-up 1). In a second approach, spores were consecutively harvested in different lots per distinct mushroom sample at distinct time points of incubation. Counted spore numbers per harvest points were added together in order to obtain total spore numbers for different lengths of incubation of a given mushroom (Experimental Set-up 2).

The standard experimental set up (caps or pieces of caps laid upside down onto wet paper with plastic lids above) was changed in some experiments by using plastic Petri-dishes without tissue paper, by using plastic Petri-dishes with a layer of Vaseline smeared onto the inner lid surfaces, by using thin glass Petri-dishes with and without a layer of Vaseline smeared onto the inner lid surfaces, and by using thin transparent plastic rings from 1 to 10 cm in height and 8.95 cm in diameter as spacers between Petri-dish bases and lids in order to adjust the relative distances between reversed mushrooms positioned on wet paper tissue in the dishes and the plastic lids above.

Evaporating dishes (9 cm Ø 4.6 cm in height) with plastic lids above were used in experiments with gasteroid mushrooms.

Germination tests

Spore suspensions (50 µl) as harvested were plated onto 2% MEA agar (20 g malt extract, 10 g agar) or LB agar (5 g yeast extract, 10 g tryptone, 5 g NaCl, 1 ml 1 N NaOH, 10 g agar) and incubated at 25 °C for up to 15 days. Mixtures of antibiotics (AB) were added to media as needed (end-concentrations: ampicillin 100 µg/ml, kanamycin 50 µg/ml, streptomycin 100 µg/ml, tetracycline 20 µg/ml, chloramphenicol 20 µg/ml). Plates were checked on daily basis for growth and nature of microbes. Where possible, colonies grown on a plate were counted.

Collection of basidiospores against gravity in plastic lids of Petri-dishes

Effects of humidity

Spores and droplets in plastic lids

Under standard incubation conditions (mushrooms laid upside-down on wet paper tissue in dishes covered with plastic lids), fine droplets usually developed overnight in the zones of the lids directly above the incubated reversed caps (Fig. 1a–d). Observations under the binocular revealed spores to be present in the droplets or, most common, spread at the surfaces of the droplets (Fig. 1e–h). Droplets with spores tended to spread somewhat irregularly flat over the plastic surfaces (Fig. 1) which suggests that they may contain some kind of surfactants. Regularly, patterns of lamellae were reflected in the lids due to the preferred positions and sizes of droplet formation. For species with dark spores, this was further visibly emphasized by their colour (Fig. 1a–f). The clear patterns of lamellae seen printed in the lids imply that spores fly straight up from their place of release toward the plastic lids where they collect with the growing droplets. Droplets however evaporated very fast upon opening of dishes. A clearly visible film of dried material was regularly left behind on the plastic surfaces (not further shown).

Spores were initially collected from the lids in 200 µl sterile water. Spores did however not easily transmit into the water but showed an affinity to stick to the plastic lid of a Petri-dish. Quick wiping with the plastic tip of a micro-pipette was required to transfer the spores into the liquid. Spores tended to clump and quickly sink to the bottoms of sterile Eppendorf cups into which spore solutions were transferred. Addition of mild detergent such as Tween 80 can help to suspend clumped basidiospores (Dhawale and Kessler 1993; Rincón et al. 2005). Therefore in later experiments, we used 200 µl sterile 0.1% Tween 80. With the detergent, spores were easily taken up from the lids and suspended.

Species range with open hymenia tested

Caps or parts of caps of mushrooms of a broad taxonomic species range were incubated upside-down on wet paper tissue in dishes covered with plastic lids. In nearly all cases (i.e. for mushrooms of 66 distinct species), basidiospores collected in large numbers in the lids (Table 2). Fresh mushrooms with open hymenia were collected over the time of species with gills (in total 59 species from 12 different families of the Agaricales—i.e. 7 species from the Agaricaceae, 2 species from the Amanitaceae, 2 species from the Bolbitiaceae, 4 species from the Inocybaceae, 3 species from the Hygrophoraceae, 3 species from the Marasmiaceae, 1 species from the Mycenaceae, 2 species from the Physalacriaceae, 1 species from the Plutaceae, 25 species from the Psathyrellaceae, 5 species from the Strophariaceae, and 3 species from the Tricholomataceae; 1 species from the Repetobasidiaceae, Hymenochaetales; 1 species from the Russulaceae, Russulales), species with gill-like ridges (1 species from the Hygrophoropsiceae, Boletales) and pseudolamellae (1 species from the Schizophyllaceae, Agaricales), and species with pores (in total 7 species from the Boletales, i.e. 6 species from the Boletaceae and 1 species from the Sulliaceae; 1 species from the Polyporaceae, Polyporales). Nearly all tested species are characterized by four-spored basidia but Agaricus bitorquis, Agaricus subperonatus, L. birnbaumii and Suillellus queletii which can have both heterokaryotic and homokaryotic spores in their caps by mixed formation of bi- and four-spored basidia (Breitenbach and Kränzlin 1991, 1995). The only four exceptions of fleshy mushrooms which failed in spore collection in our experiments were single mature individuals of Hygrocybe virginea and Hygrophorus eburneus (both Hygrophoraceae), Lepista saeva (Tricholomataceae) and Rickenella fibula (Repetobasidiaceae). Since other species of the same or a closely related family gave spores in the lids (Table 2), these four failures did not relate to any specific taxonomic position of mushrooms but possibly to that spore shedding ended by fruiting body age (Haard and Kramer 1970; Li 2005; Saar and Parmasto 2014; not further analysed).

Spore solutions from mushrooms incubated upside-down overnight in Petri-dishes typically contained between about 10⁴ up to in highest cases >10⁷ total spores (compare Table 2). Spore yields obtained from different mushroom samples of a same species were usually very similar, even when harvested and incubated at different days (details not further shown but see the averaged data in Table 2 and compare the data for individual species also with those from specific experiments presented in Tables 1, 3; Figs. 2, 3). This suggests in coincidence with earlier reports on spore releases of different species (Fischer and Money 2009; Saar and Salm 2014) that species-specific parameters determine the frequency of spore release and the actual spore capture in the plastic lids. However, spore yields did not plainly depend on a single simple parameter such as cap size, spore sizes, structures of hymenia (lamellae, ridges or pores) (Table 2), or gill numbers per cap and pore diameters (see the cited field guides for the individual parameters of species for comparison with the spore collection data in Table 2). Cap age and speed of younger cap maturation, lengths and mode (consecutive or synchronous) of spore production and maturation periods (Kües and Navarro-Gonzaléz 2015; Halbwachs and Bässler 2015) might also be needed to be considered on individual species level as parameters of potential influence on spore harvests.

Species	No of mushrooms per situation	Part used	No of samples	Spore harvests
				Plastic lid		Glass lid
				Plain	With Vaseline	Plain	With Vaseline
Coprinellus micaceus	2	1/2	3	5.1 ± 3.0 × 105	2.1 ± 1.0 × 105	6.0 ± 3.3	7.3 ± 5.3
Coprinellus domesticus	3	1	3	4.9 ± 1.4 × 106	1.6 ± 0.5 × 106	11.0 ± 6.5	20.0 ± 13.1
Coprinellus disseminatus	5	1	5	2.1 ± 0.9 × 105	4.5 ± 1.1 × 105	11.7 ± 9.4	24.0 ± 19.6
Coprinopsis atramentaria	1	1/2	2	5.9 ± 0.4 × 106	4.1 ± 0.4 × 106	0	0
Coprinus comatus	1	1/8	2	9.2 ± 0.4 × 106	5.7 ± 0.5 × 106	0	0
Hypholoma fasciculare	3	1	3	1.0 ± 0.5 × 105	0.9 ± 0.5 × 105	0	0
Psathyrella candolleana	1	1	1	9.4 × 105	7.6 × 105	0	0

Effects of mushroom conditions on spore harvests

We collected mostly fresh fleshy mushrooms of Agarics (Table 2) and many were still young, in the stage of opening or close to be fully opened. During overnight incubation in reversed orientation, younger caps of most of the fleshy species further opened and cap diameters (measured at the time of harvest from nature, see Table 2) tended to further extend by stretching out the umbrellas. Of the lamellate species, the single fruiting bodies of Agaricus campestris, Agaricus subfloccosus, Hygrophorus olivaceoalbus, Inocybe fraudans, Marasmius oreades, and Psathyrella spadiceogrisea, the two mushrooms of Pluteus spec., all four of Marasmius wynnei and all six of Psathyrella candolleana were already fully open at their harvest. There were no problems with any of the younger and the here listed mature mushrooms to obtain high numbers of spores transferred to the plastic lids positioned above (Table 2).

Notably, also younger (P. conopilus), mature (the single fruiting bodies of Agaricus augustus, Pholiota spec. and Stropharia caerulea; one mushroom each of P. candolleana and S. commune, two mushrooms each of Panaeolus ater and Psathyrella atrolaminata, and a series of mushrooms of C. domesticus) or aging mushrooms (Conocybe tenera, one mushroom each of Agrocybe dura, Marasmius cohaerens and Panaeolus cinctulus) which were to different extend desiccated were successfully appointed in collecting spores in numbers of 10⁴–10⁶ in plastic lids by reversed incubation on wet paper tissue (Table 2; Fig. 2b). The shrivelled mushrooms refreshed in shape by taking up humidity from the wet paper tissue. For A. dura, we had an older dry fruiting body and young fresh mushrooms. Spore harvests were similar (2.6 × 10⁶ spores versus from 2.0 to 3.5 × 10⁶ spores). Spore numbers for dehydrated mushrooms of C. domesticus were only somewhat reduced (0.9 ± 0.1 × 10⁶; Fig. 2b) as compared to most mushrooms harvested in fresh stage (4.1 ± 1.8 1 × 10⁶; Table 2). Moreover, mushrooms of the durable species S. commune which were dried on purpose at RT for 15 days gave still considerable numbers of spores in lids (5.6 ± 0.8 × 10⁴; n = 3; tested with the water agar system) over the 18 h of reversed incubation although these were 100× reduced as compared to mushrooms which were used directly at the day of harvest (Table 1). S. commune in active phases produces continuously new spores (Kües and Navarro-Gonzaléz 2015) by which fresh mushrooms might distinguish from revived specimens that will require time for full physiological recovery.

Further of importance, also mushrooms infested with small animals (Fig. 4) can be used in collecting spores in lids. Of the lamellate mushrooms mentioned above, Pluteus spec. for example carried small slugs (Fig. 4a, b), on P. spadiceogrisea and A. campestris were rove beetles (Fig. 4d) and red mites and internally some larvae (Fig. 4g, h), while the ones of A. campestris (Fig. 4h), A. subperonatus (Fig. 4i–k) and M. wynneae (Fig. 4e, f), one of six of P. cinctulus (Fig. 4l, m) and one of two mushrooms of Pholiotina vestita (Fig. 4c) had grubs. Particularly the fleshy pilei of the A. campestris (Fig. 4h) and A. subperonatus fruiting bodies (Fig. 4i–k) and also all mushrooms of the Boletales (Fig. 4n and not shown) were much infested with many grubs. Insect larvae had eaten tunnels into the pilei of the S. queletii (Fig. 4n) and the other Boletales’ mushrooms. However, there was still much surface area with intact pores (Fig. 4n and not shown) in order to obtain bulks of basidiospores in plastic lids above (Table 2). Only for an aged, slug-eroded and fully grub-populated decaying Xerocomellus chrysenteron fruiting body, it was not possible to obtain spores from. Where we had infested and animal-free fruiting bodies, harvested spore numbers were still similar (infested decaying C. domesticus fruiting body: 0.9 × 10⁶ spores, animal-free mushrooms: from 1.3 to 6.9 × 10⁶ spores; infested P. cinctulus fruiting body: 0.9 × 10⁶ spores, animal-free mushrooms: from 2.9 to 5.4 × 10⁶ spores).

Spore release over the time

Mushrooms started quickly to propel off spores when incubated upside-down on wet tissues in closed dishes (Fig. 2). Already after 2 h of incubation, considerable amounts of spores (>10⁴–10⁵ for most species or for somes species even 10⁶) could be harvested from mature fruiting bodies in plastic lids. Fast spore release continued for a few hours (4–6 h) but the speed of spore release usually decreased for several of the species with time of incubation to eventually level off to a maximum amount of spores which possibly can be discharged by a single specimen under the experimental conditions applied.

In many cases, we used in our experiments fleshy mushrooms of limited life-time such as of the ephemeral inkcaps (species of Coprinellus, Coprinopsis, Coprinus) that after a transitory period of ballistospore ejection release a majority of their spores in liquid droplets by autolysing their caps (McLaughlin et al. 1985; Kües 2000; Redhead et al. 2001; Nagy et al. 2013). The time of spore release by ballistospore ejection up to onset of cap autolysis was always sufficient to obtain high spore numbers in lids of plastic Petri-dishes, in amounts of >10⁵ to >10⁶ (Tables 1, 2, 3; Figs. 2, 3). In case of C. cinerea fruiting bodies from laboratory cultures, spore release was delayed by 4 h from the start of incubation of young opening caps with still pale gills (Fig. 2e). This time coincided well with the known time schedule of basidiospore maturation with corresponding black gill staining after light-induced synchronized karyogamy in the basidia (Kües and Navarro-Gonzaléz 2015).

Spore attachment to lids of different material

In preliminary experiments, mushrooms of S. commune and of C. atramentaria were incubated upside-down in dishes with plastic lids or in dishes with glass lids. Spore collected overnight in plastic lids whereas spores were not present in glass lids (data not shown). The results suggested that an electrostatic disposition possibly helped to attach ejected spores to the surface of the plastic lids.

We smeared sticky Vaseline onto glass and plastic lids of Petri-dishes to support attachment of spores to the surfaces. Experiments with parallel sets of overnight upside-down incubated mushrooms or parts of mushrooms of Hypholoma fasciculare, P. candolleana and five different inkcap species showed that Vaseline in plastic lids had no incisive negative effect on spore yields (Table 3; Fig. 1e–h). Spores stuck well in droplets to the Vaseline (Fig. 1f, h) although it was harder to harvest them from Vaseline and bring them into solution than without. However, Vaseline did not lead to considerably increased numbers of spores that attached to glass lids. With and without Vaseline, there were always either no or only negligible few spores in glass lids (Table 3). We conclude from the experiments that different affinities between the glass and plastic surfaces of tying spores is not the primary cause for the unequal spore harvests from lids of different material but a distinct effect of the plastic on upward spore transfer. Likely, static electricity of the plastic (Woodland and Ziegler 1951; Kuo 2015) will force spores to fly upward onto the lids where the same forces will then support attachment of the spores to the plastic surface.

Spore transfer to plastic lids over different distances

The data in Table 1 implicate for several species that not all basidiospores released from reversed mushrooms are attracted to plastic lids. Experiments in which the distance between the upside-down laid mushrooms and the plastic lids above for several species were varied further support an influence by spore properties on spore yields in the lids. For all species, numbers of spores attached to the lids decreased in linear trends on the log scale with increasing distance to the reversed mushrooms (Fig. 3). Roughly, three groups in line steepness might be distinguished (Fig. 3a, d, j–l, steepest: C. comatus, A. solidipes, Lepista nuda, P. squarrosa, S. commune; Fig. 3b, e, g, h, medium: C. cinerea, C. atramentaria, C. domesticus, Tubaria furfuracea; Fig. 3c, f, i, flattest: C. truncorum, C. micaceus, Kuehneromyces mutabilis). Break-offs of all lines (lowest distances where no spores detected in lids) were abrupt. Between species, break-offs occurred at different heights, following different orders of magnitudes (ranging from <10⁴ up to nearly 10⁶) of absolute spore numbers at the last positive height. Absolute numbers of spores collected in lids at low distances did not correspond throughout to the possible longest distance over which spores of a species were found to be attracted to plastic lids (Fig. 3). C. atramentaria as a species with highest spore numbers in lids at short distance (>10⁷ at 1 cm distance) also yielded reasonable amounts of spores in lids at highest distances (>10⁵ at 7 cm distance; Fig. 3e), followed by Coprinellus truncorum and C. micaceus with both ca. 4 × 10⁵ spores in lids at 1 cm distance and ca. 5 to 7 × 10⁴ spores at 6 cm distance (Fig. 3c, f). In all other cases, maximum heights of spore detection varied between 3 cm and 5 cm distance. Spore sizes (lengths, see Table 1) or volumes (as calculated for an ellipsoid V = 4/3 π × ½ length × 2 × ½ width; data not shown) did also not correlate with maximum heights at which spores were attracted to plastic lids (Fig. 3). Attraction between higher up-flying spores and the plastic lids might be expected to be stronger than between less far up-flying spores and the lids. Basidiospores do also have electric charges (Buller 1909, 1922; Gregory 1957; Swinbank et al. 1964; Webster et al. 1988; Saar 2013; Saar and Parmasto 2014; Saar and Salm 2014). Individual differences in charging of spores remain as explanation for the observations.

Capturing spores from gasteroid basidiomycetes

We concluded before that spore collection in plastic lids positioned above mushrooms with open hymenium depended on their forcible ballistospore discharge mechanism mediated at high humidity by the fusion of Buller’s drops with liquid films at the adaxial sides of the spores. Gasteroid species have lost this active basidiospore discharge mechanism, produce their basidiospores within closed fruiting bodies and disperse them from openings or cracks passively with wind or through pushing the spore sacs by rain drops (Hibbett et al. 1997; Wilson et al. 2011; Kües and Navarro-Gonzaléz 2015). Accordingly, basidiospores did not accumulate during overnight windstill incubation in evaporating dishes (18–24 h) in the plastic lids above mature puffballs (Lycoperdon perlatum, Agaricaceae) which were either already cracked or of which the inner gleba with the basidiospores was opened by cutting the fruiting body into two equal halves. Similarly, there were no spores in lids after 18 h incubation when we positioned two different mature earth-stars (Geastrum rufescens and Geastrum striatum; Geastraceae, Geastrales) with an ostiole as a natural opening and unrolled segments of the exoperidium into windstill evaporating dishes covered by plastic lids. However, when we hit 2× with caution the spore sacs from the side with a glass rod through the spouts of the evaporating dishes, clouds of spores escaped through the ostioles from the spore sacs into the air and attached to the plastic lids above (not further shown). We harvested and counted in the lids then 6.9 × 10⁶ basidiospores of G. rufescens (plating proofed them to be contamination free) and 4.4 × 10⁶ for G. striatum. We conclude that the spores are moved into the air is decisive for spore attraction to the plastic lids but not the particular initial mode of the spore release.

Spore plating

The reliable yields of spores from upside-down positioned wild mushrooms in the plastic lids of Petri-dishes evoked the further idea to test whether just basidiospores were released onto the lids. First, 50 µl aliquots of spore solutions from individual mushrooms from the Experimental Set-up 1 in Table 1 were analysed on MEA media. Spore solutions from mushrooms incubated in Petri-dishes in natural direction gave rise to massive bacterial and also numerous fungal contaminations, in contrast to aliquots from spore suspensions which were harvested from plastic lids from upside-down incubations of mushrooms (Table 4). Generally, sizeable bacterial colonies appeared on media after 1 day at 25 °C incubation, colonies of yeasts and molds after 2 days and basidiospore germlings visible to the naked eye for all tested mushrooms after 3 days, respectively. Both bacterial and fungal contaminants (molds, mostly Ascomycetes) hindered by overgrowth other organisms in growth, including germination of the basidiospores and growth of germlings of the respective mushroom plated (Table 4). Addition of antibiotics suppressed bacterial growth but fast growing molds present in the samples of classic spore prints still overgrow the basidiospores (Table 4; Fig. 5a). Colonies from germinated basidiospores of spore suspensions from classical prints among contaminations were therefore only observed in the cases of C. domesticus and S. commune but at much lower frequency (≥1000 fold less) than on plates onto which samples of basidiospores were plated from harvests from plastic lids located above upside-down incubated mushrooms (Table 4). In absence of contaminants, germination of basidiospores from samples harvested from plastic lids was unhindered for all mushrooms of the four species tested. There were no bacterial contaminations and fungal contamination occurred only in one instance (=10% of all tested cases) with just 3 yeast colonies (Table 4; Fig. 5b). While the germination rates of basidiospores were different between the species used (Table 4), possible effects by densities of the own basidiospores in the solutions were not further tested.

Fruiting body	Classic spore print (mushroom upside-top)
	Basidiospores in 50 µl	Colonies on plate/50 µl spore solution plated
		MEA			MEA + AB
		Basidiomycete	Contaminationa		Basidiomycete	Contaminationa
			Bacteria	Other fungi		Bacteria	Other fungi
Coprinellus domesticus	1.6 × 107	68	Uncountable	2 yeasts, 34 molds	51	–	122 yeasts, 14 molds
Panaeolus cinctulus	8.0 × 106	–	5.4 × 103	3 yeasts, 38 molds	–	–	8 yeasts, 14 molds
Schizophyllum commune	2.1 × 107	2563	Uncountable	34 yeasts, 4 molds	2326	–	544 yeasts, 8 molds
Tubaria hiemalis (1st mushroom)	1.9 × 104	–	1.7 × 105	14 yeasts, 6 molds	–	–	42 yeasts, 10 molds
T. hiemalis (2nd mushroom)	2.4 × 104	–	4.3 × 105	6 yeasts, 5 molds	–	–	64 yeasts, 21 molds

	Spore print in plastic lid (mushroom upside-down)
	Basidiospores in 50 µl	Colonies in plate/50 µl spore solution plated
		MEA			MEA + AB
		Basidiomycete	Contamination		Basidiomycete	Contamination
			Bacteria	Other fungi		Bacteria	Fungi
C. domesticus	1.2 × 106	4660 (0.33% germination)	–	–	4852 (0.34% germination)	–	–
P. cinctulus	9.0 × 105	1352 (0.15% germination)	–	–	1280 (0.14% germination)	–	–
S. commune	1.8 × 106	Uncountable	–	–	Uncountable	–	–
T. hiemalis (1st mushroom)	7.8 × 103	2458 (31.5% germination)	–	–	2873 (36.8% germination)	–	–
T. hiemalis (2nd mushroom)	9.0 × 103	2213 (24.5% germination)	–	–	2437 (27.0% germination)	–	3 yeasts

^aNumbers of fungi in different media did not clearly correspond to each other since fast bacterial growth suppressed fungal growth and fast growing molds suppressed that of other molds and yeasts

Second, spore suspensions from classical prints of C. domesticus and P. papilionaceus mushrooms fixed to dishes by agar during downward spore shedding were compared on LB medium at 25 °C with spore suspensions obtained by harvests from plastic lids from agar-fixed mushrooms incubated upside-down (solutions from the Experimental Set-up 2 in Table 1). In case of the classical C. domesticus spore print (5.3 × 10⁶ spores/50 µl), >360 large slimy and blurred bacterial colonies grew over the surface of the plate and five yeast colonies on medium with antibiotics while the plates with spore suspensions from the upside-down incubated fruiting body (6.0 × 10⁵ spores/50 µl) were all free of any contaminations (but also of germinated basidiospores; not further shown). Judging by colony morphologies, multiple bacterial species (>10) grew in high density of a plated basidiospore suspension from the classical spore print of P. papilionaceus (8.8 × 10⁵ spores/50 µl) and still three types of bacteria (338, 15 and 3 colonies, respectively) when antibiotics were added to the growth medium. This contrasted the situation with the spore solution from the upside-down incubated mushroom (1.0 × 10⁵ spores/50 µl) where no bacteria were found on LB without or with antibiotics while germination of basidiospores at 0.72% frequency was observed (not further shown).

From the time course experiment presented in Fig. 2, we had each two samples for C. domesticus (Fig. 2b), C. micaceus (Fig. 2b) and P. cinctulus (Fig. 2a) per tested time point of spore release into lids of plastic Petri-dishes. We tested also these for presence of contaminations by cultivation on MEA at 25 °C. In case of C. domesticus 26 of 30 basidiospore samples tested (87%) were free of contaminants, in case of C. micaceus 22 of 30 samples tested (73%), and in case of P. cinctulus 30 of 40 samples tested (75%). When contaminants were observed (in 22% of all these cases together), these were always bacteria (usually 1 or 2 types, rarely 3 types of bacteria) while the number of bacteria per 50 µl plated spore suspensions differed between 3 and 407 for C. domesticus, 3 and 14,400 for C. micaceus and between 2 and >10⁶ for P. cinctulus (Fig. 6). There was a tendency in likelihood for samples obtained from longer mushroom incubation to contain contaminations but there was no continuous increase in bacterial numbers over the different samples with length of mushroom incubation applied for spore release. Moreover, samples from two different halves of a same mushroom often differed in that bacteria were observed in only one of the two (Fig. 6). Furthermore, spore samples (50 µl) of the each four analysed time periods (5, 10, 15, 20 h) of spore release of A. strobiliformis (Fig. 2b), A. subperonatus (Fig. 2a), C. piperatus (Fig. 2b), M. wynneae (Fig. 2a), and Suillus spec. (Fig. 2b) were all free of bacteria but the 15 h sample of C. piperatus (1 bacterial colony), the 20 h sample of M. wynneae (1124 bacterial colonies) and the 20 h sample of Suillus spec. (2 yeast colonies). Thus, also in these series of samples, when contaminations were found (in 15% of all cases; 10% bacterial contaminations; 5% fungal contaminations) they came from longer mushroom incubations appointed for spore shedding.

Finally, spore solutions (50 µl) from 18 or 20 h incubation of mushrooms in upside-down position of a broader range of 19 randomly collected species (from Table 2) were tested on MEA. No contaminants were found in spore suspensions of individual fruiting bodies or of parts of fruiting bodies of A. augustus, A. bitorquis, A. campestris (grub infested; Fig. 4g, h), A. subfloccosus, A. subperonatus (grub infested; Fig. 4i–k), Amanita excelsa, Hygrocybe conica, L. sulphureus, L. birnbaumii, Pholiota spec., P. vestita (of two tested, one was grub infested; Fig. 4c), Psathyrella spadiceogrisea (with beetle; Fig. 4d), and X. chrysenteron (grub infested). Contaminations were found in one of two tested A. dura spore solutions (203 colonies, 1 type of bacteria), in one of the solutions from two tested grub-infested Boletus luridus fruiting bodies (20 colonies of yeasts), and in spore solutions of H. olivaceoalbus (59 bacteria, 1 type), Inocybe erubescens (3 yeast colonies), and grub-infested mushrooms of Boletus rhodoxanthus (312 colonies, 2 types of bacteria) and S. queletii (2128 colonies, 3 types of bacteria); i.e., contaminations in spore solutions in this series of platings were discovered in solutions of 27.3% of all tested mushrooms (i.e. in 20.7% of all individual samples tested) with 18.2% (13.8%) bacterial infections and 9.1% (6.9%) fungal contaminations. As a further important aspect, it should be recalled that in total 9 of these 22 mushrooms tested (Fig. 4d and not further shown) were infested with animals while only 3 spore solutions of these were contaminated.

Considering all plating test series together, 19 mushrooms infested with animals were analysed (from 15 different species; 28 different samples). Spore solutions of only 7 of these (25.0% in total; 17.9% with bacteria, 7.1% with fungi) were found contaminated.

Driving forces for spore flight

If only the catapulting energy and gravity will be the acting driving forces for the spores to move upon release, accumulation of basidiospores against gravity in the lids of plastic Petri-dishes would not be possible. The energy of catapulted basidiospores is quickly used up during the propelling into the free airspace by the braking effect through the viscosity of the air (Turner and Webster 1991; Pringle et al. 2005; Stolze-Rybczynski et al. 2009; Fischer et al. 2010a, b). Once fully braked, the spores should then drift downward by action of gravity, which in case of upside-down positioned mushrooms would be down into the free gaps between hymenia. This reasonable prospect made us wondering what in our experiments might be the reason for about 10% to up to 100% of the basidiospores from a mushroom to instead fly up and attach to the plastic Petri-dish lids (Table 1).

The air space in the closed Petri-dishes can be expected to be motionless. Concerted release of basidiospores by a mushroom and evaporate cooling of the air surrounding the cap can principally help that the spores are whirled up in clouds by created airflows (Buller 1934; Deering et al. 2001; Dressaire et al. 2015, 2016). However, the clear lamellar patterns of mushrooms reproduced in our experiments by the spore repositories in the plastic lids speak against any generation of influential eddies in the closer head space of the mushrooms. Our investigations revealed that electrostatic charges by plastic are the likely reason for spores to move straight up against gravity and also for them to attach to the attracting plastic (Table 1). However, charges innate to the spores should also have their part in this process (Fig. 3). Basidiospores when ballistically discarded from mushrooms are indeed electrically charged. Charges of endogenous origin are possibly connected to the mass transfer when Buller’s drop suddenly fuses with the liquid film on the adaxial side of a spore. In addition, airflow will create charges on spores by triboelectric effects (Buller 1909; Webster et al. 1988; Saar and Parmasto 2014; Saar and Salm 2014) which could give an extra stimulus to the spores to leave the caps against gravity by electrostatic attraction to plastic lids.

In species-specific manner, spores of an individual mushroom can differ in type of charge (+ or −) and in strength of charging. The charges cause in horizontal electric fields that spores shed from mushrooms will drift away to one side. In many species, there are populations of positively and of negatively charged spores, while spores in some species are unipolar-positively and in some others unipolar-negatively charged (Buller 1922; Gregory 1957; Webster et al. 1988; Saar 2013; Saar and Parmasto 2014; Saar and Salm 2014). Of the species that we tested in this study (Table 2), absolute charges of basidiospores have been estimated before for A. campestris in different experimental series between 1.55/0.39/0.39 and 3.10/3.11/2.16 × 10⁻¹⁷ C (mean 2.23/1.28/0.93 × 10⁻¹⁷ C), for C. micaceus between 0.85 and 4.22 × 10⁻¹⁷ C (mean 2.14 × 10⁻¹⁷ C), for L. nuda 1.88 × 10⁻¹⁷ C, for M. oreades between 1.25 and 10.12 × 10⁻¹⁷ C (mean 4.37 × 10⁻¹⁷ C), for two Pholiota spec. strains between 1.65 and 2.03 × 10⁻¹⁷ C, and for S. commune between 1.70 and 9.55 × 10⁻¹⁷ C (mean 4.10 × 10⁻¹⁷ C), (Webster et al. 1988; Saar and Salm 2014). Populations of basidiospores of L. nuda were found to be 97% negatively charged, whereas populations of A. campestris and Pholiota species were bipolar with more numerous positively charged spores and populations of C. micaceus bipolar with more negatively-charged spores (Buller 1909; Gregory 1957; Saar 2013; Saar and Salm 2014).

Depending on individual spore charges and on relative distances (Fig. 3), electrostatic charged plastic lids of the Petri-dishes may thus differentially attract the also loaded spores. This can then explain why in our experiments mostly only parts of the total spores released from a fruiting body are directed toward the plastic lids positioned above mushrooms (Table 1) and why with higher distances between lids and mushrooms the number of spores attached to the lids decreased in logarithmic fashion (Fig. 3). Overall charges of spores correlate little with spore sizes and actual spore charges are also independent on spore emission rates from mushrooms (Saar and Salm 2014). This is also reflected in our results on different species as shown in Fig. 3. Spore sizes have in contrast been shown to influence sizes of Buller’s drops, the spore velocity upon ejection from the sterigmata, the length of the move vertically through a mushroom airspace before being braked (Stolze-Rybczynski et al. 2009; Fischer et al. 2010a), and the distance of spore deposition from the source (Norros et al. 2014), parameters which appear all be not of primary relevance to the electrostatic attraction and attachment of the spores to plastic lids reported here.

Incidence of basidiospore charging in Agaricomycetes

In this study, basidiospores of in total 66 species with open hymenia and ballistospore propulsion mechanism (from 36 genera, 19 families and 5 orders) did accumulate against gravity in plastic lids and we failed only for four species with each one mushroom tested (H. virginea, H. eburnus, L. saeva, R. fibula) to proof such effect. Charging of ballistospores appears thus to be a widely distributed property in the Agaricomycetes. This confirms former observations by Saar and Salm (2014). These authors concluded from own and literature data for 43 species, 33 genera, 23–25 families and 9 orders of the Agaricomycetes that their ballistospores exhibit electric loads. While we used in our studies in the majority species of the Agaricales and to less extend species from the Boletales and other orders (Table 2), Saar and colleagues analysed the electric properties of larger species ranges of Polyporales and Boletales with poroid hymenia, in addition to species from the Agaricales, Russulales and others (Saar 2013; Saar and Parmasto 2014; Saar and Salm 2014). Considering shared species between the different studies, evidence for ballistospore electric charging is now available for over 100 different species (i.e. 103 or 104 species) of the Agaricomycetes.

As we have shown here, charging is not only confined to ballistospores of Agaricomycetes. The spores of earthstars which lost the mechanism of ballistospory (Hibbett et al. 1997; Wilson et al. 2011) were also attracted by plastic lids once they were manually pushed into the air. They attached even stronger to the plastic lids than the spores of other Agaricomycetes. When trying to brute-force them into solution, they repeatedly formed a dense layer as a cover over the surface of a sphere of 200 µl water. The water ball with the spores on the outside repeatedly burst to slip away from underneath the spore layer which remained then as dry dark brown spot (about 7 mm in Ø) of densely packed spores strongly attached to the plastic surface of the lid (our unpublished observations).

Microbial contaminations in spore solutions

We have demonstrated that basidiospore transfer from upside-down incubated mushrooms with open hymenia to the plastic lids depends on an active ballistospore discharge mechanism (Table 1). This suggested that producing spore prints from up-flying spores would be more selective to the basidiospores than the usual spore prints collected underneath wild mushrooms which tend to be mixed with fallen spores and cells of other microbes. This assumption was tested by plating spore solutions from classical prints and spore solutions collected from plastic lids positioned above upside-down incubated mushrooms (Table 4; Figs. 5, 6). In contrast to conventional spore prints, presence of yeasts or spores from other filamentous fungi which need animal vectors or air flow for their movements was rarely detected in plating tests of spore solutions which were collected from plastic lids positioned above reversed incubated mushrooms (at maximum 10% of samples were contaminated). Bacteria were in somewhat higher frequency present (in up to 22% of total samples of an individual test series), possibly because they may at times be flying directly with a basidiospore. However, there are possible measures against bacteria through addition of suitable antibiotics to the growth media (Fig. 5; Table 4). Further, accumulation of larger numbers of bacterial cells can be precluded by shorter incubation times applied for basidiospore collection (Fig. 6). Already 2–4 h of incubation are sufficient to collect 10⁴–10⁵ spores from mushrooms in the plastic lids (Fig. 2) with no or exceptionally a few bacteria being present in the spore solutions (Fig. 6). Spore ejection rates in our study (Fig. 2; see Table 1 for total ejection figures) correlate with ratios of ballistospore ejection/h from mushrooms reported by Buller (1909, 1922) and to ratios which can be calculated from the spore emission data/s × cm² compiled in Saar and Salm (2014).

Basidiospores secrete hygroscopic hexoses and alcohols (mannitol) localized at their hilar appendices for the assembly of Buller’s drops and at their adaxial shallow dents as hygroscopic compounds for the formation of liquid films, both of which are required for rapid spore catapulting upon their fusion (Goates and Hoffmann 1986; Webster et al. 1989, 1995; Turner and Webster 1995). These organic metabolites together with inorganic ions like phosphate, sodium and potassium are transferred with the liquids spread over the propelled-off spores onto the lids of the Petri-dishes where they might act further hygroscopically to attract more water to the spores (Elbert et al. 2007; Hassett et al. 2015). Eventually, the metabolites might be used for growth of any microbial contaminants happened to be present in the liquid droplets in which basidiospores amplify on the plastic lids during mushroom incubation (Fig. 1). Such microbial growth effect can explain the explosive sudden increase in bacterial cells which was observed more often in spore samples collected after longer incubation times of mushrooms (Fig. 6).

Accordingly, earlier spore harvests can reduce the chance for unwanted bacterial transfer and the time for possible proliferation of contaminants. For later use after a time of storage, drying off the liquid and storing the basidiospores under dry conditions on the sterile plastic lids is also a possible measure of avoidance. Earlier harvest times furthermore reduce the danger of undetected small animals to creep out of mushrooms and to crawl over and contaminate with other microbes the spore collections assembled over the time above the mushrooms on the plastic lids. Such crawling of initially overlooked insect larvae over the spore prints in the plastic lids has indeed been noticed by us on two occasions (not further documented).

Spore charges in applications

Charging of fungal spores and any potentially connected functions (e.g. in support for spores to serve as nuclei for raindrops; Elbert et al. 2007; Hassett et al. 2015) are understudied and generally little understood (Webster et al. 1988; Wargenau et al. 2011, 2013; Saar 2013; Saar and Parmasto 2014; Saar and Salm 2014). In line, practical use of charging of spores and spore behaviour in electric fields has so far only seldom been made of. However, guarding of bookshelves with an electric field screen has been tested to successfully protect old valuable books in library stack rooms against mold infection by airborne spores (Takikawa et al. 2014) and electrostatic dust collectors were applied in French archives to study their fungal allergenic potentials (Roussel et al. 2012). Spores in buildings and rooms distribute unevenly, among as factors behind by being influenced by electric and magnetic devices (Anaya et al. 2015, 2016). Knowledge on fungal spore distributions in buildings as in outside environments can come through application of electrostatic dust collectors (Normand et al. 2016) and electrostatic precipitators might be used (Han et al. 2011). An electrostatic nursery shelter and an ozone-generative electrostatic spore precipitator were reported to protect tomato plants in hydroponic culture and in open-window greenhouses against fungal pathogens (Shimizu et al. 2007; Kakutani et al. 2012; Takikawa et al. 2016) and an electrostatic spore collector has been applied to collect conidia of the barley pathogen Blumeria graminis (Moriura et al. 2006). While our experimental set-up here is simple and easy to apply by using electrostatic features intrinsic to the plastic lids of Petri-dishes to selectively collect basidiospores from mushrooms, our observations offer the possibility to also develop technically more sophisticated devices for basidiospore collection.