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Phytologia (Dec 20, 2024) 106(4) 51
Sommerstorffia pugioniformis sp. nov: A new species of rotifer-trapping oomycete
discovered using a novel baiting technique
Paul G. Davison
Department of Biology, University of North Alabama, Florence, AL 35632, USA, pgdavison@una.edu
Jeffery M. Ray
Department of Biology, University of North Alabama, Florence, AL 35632, USA, jmray1@una.edu
Carrie Ann Davison
Department of Genetics, Yale University, New Haven, CT 06510, USA, carrieann.davison@yale.edu
ABSTRACT
The genus Sommerstorffia (Oomycota, Saprolegniales, Verrucalvaceae) is a predator of freshwater
rotifers last recorded in North America sixty years ago. Here we describe a novel baiting technique for
Sommerstorffia that relies on the easily cultured rotifer Lecane inermis and plastic coverslips as a
transparent substrate for Sommerstorffia’s attachment. The efficacy of the baiting technique is supported
by our multiple records of Sommerstorffia spinosa across seven U.S. states. With this technique, we
discovered sporeling morphology quite different from S. spinosa leading us to propose Sommerstorffia
pugioniformis as a new species. Molecular analysis based on COI and ITS regions placed the new species
and S. spinosa within the Verrucalvaceae clade. Published online www.phytologia.org Phytologia 106(4):
51-67 (December 20, 2024). ISSN 030319430.
KEY WORDS: Lecane inermis, Sommerstorffia pugioniformis, S. spinosa, baiting, oomycete, predatory
fungi, standing drop, rotifers, Verrucalvaceae.
_____________________________________________________________________________________
Several lineages of fungi and fungal-like organisms are known to parasitize or prey on rotifers
(Barron 2004; Glockling et al. 2014). Within the monophyletic Verrucalvaceae (Oomycota, Saprolegniales)
three genera include species parasitic/predatory on rotifers. The taxonomically diverse genus Aphanomyces,
best known for several economically important species parasitic on plants and animals (Becking et al.
2022), includes two questionable species, A. americanus and A. hydatinae, that are parasitic on rotifers but
are not known beyond their original descriptions (Scott 1961; Molloy et al. 2014). More recently, a new
genus Aquastella was described to accommodate two species that are obligate rotifer parasites, A. acicularis
and A. attenuata, known only from planktonic rotifer hosts collected over a seven-year period in Brooktrout
Lake, Adirondack Park, New York State (Molloy et al. 2014). Finally, Sommerstorffia, with only S. spinosa
Arnautov (1923a), is a globally widespread predator of rotifers.
Sommerstorffia spinosa is a predator of common, periphytic, loricate rotifers in freshwater habitats
(Glockling et al. 2014). Sommerstorffia’s structure and development was described in detail by Karling
(1952) who made a prolonged study of a culture derived from an Alaskan alpine tundra stream. Even to a
nonspecialist, Sommerstorffia is quite recognizable based on the distinctive stiff hyphal branches that end
in a predaceous peg. Sporelings that form from encysted zoospores also bear a predaceous peg. Rotifers
grazing for food bite the predaceous pegs and thereby become trapped by the mouth by an adhesive
substance (Karling 1952; Saikawa and Hoshino 1986; Saikawa et al. 2021). A lobed zoosporangium forms
inside the body of the rotifer, and zoospores are released through an exit tube in achlyoid fashion.
The published discoveries of Sommerstorffia spinosa appear to have been made without prediction
and were typically made weeks (Karling 1952; Saikawa and Hoshino 1986) or months (Sparrow 1929;
Prowse 1954; Czeczuga et al. 2000) after initiation of gross cultures usually augmented with traditional
baits (e.g., onion skin, snake skin, plant seeds) for the recovery of various saprobic water molds.
Phytologia (Dec 20, 2024) 106(4)
52
Presumably these lengthy intervals between culture initiation and Sommerstorffia detection allowed host
rotifer populations to increase and thereby favor Sommerstorffia’s rise to detectable levels. The
inconspicuous nature of S. spinosa and its dependence on rotifers susceptible to the trapping mechanism
may be among the reasons few freshwater mycologists have reported Sommerstorffia in published field
studies of oomycetes. To our knowledge, the last known published observation from North America was
made in 1963 by Charles E. Miller from Mountain Lake Biological Station, Virginia (Miller 1965).
As a result of collecting samples for an undergraduate botany course, P. G. Davison observed
Sommerstorffia spinosa from two springs in north Alabama in 2011 and 2013. Since 2013, P. G. Davison
developed a baiting technique that allows for relatively rapid lab-based detection of Sommerstorffia in field
samples. The baiting technique relies on the easily cultured rotifer Lecane inermis (Bryce, 1892) as bait.
We applied this method to a sample from Mill River, Connecticut, which led to the discovery of
Sommerstorffia nearly identical to S. spinosa except for the unique shape of its sporelings. We describe the
novel baiting technique and our method of culture from single spore isolates below along with a description
of the proposed new species Sommerstorffia pugioniformis. Our observations provide the first documented
occurrences of S. spinosa in Alabama, Connecticut, Florida, Mississippi, North Carolina, and Tennessee.
In 2013 Alabama material of S. spinosa fixed in RNAlater was sent to Christoffel F.J. Spies and C.
André Lévesque who isolated the LSU and SSU regions (unpublished, see Glockling et al. 2014; Molloy et
al. 2014) that indicated Sommerstorffia was closely related to Aphanomyces as had been previously
suggested by Arnaudov (1923a) and Karling (1952). Based on trustworthy, though unpublished, sequence
data Sommerstorffia’s placement in the Verrucalvaceae was accepted (Beakes and Thines 2017, Rocha et
al. 2018). We generated and analyzed Cytochrome c oxidase subunit I (COI) and the internal transcribed
spacer (ITS) recommended by Robideau et al. (2011) as the complementary DNA barcodes used for
taxonomic identification of oomycetes. With this analysis, we provide the first published sequences
supporting Sommerstorffia’s placement in the Verrucalvaceae. Additionally, we provide sequence data in
support of recognizing two species of Sommerstorffia, S. spinosa and S. pugioniformis.
MATERIALS AND METHODS
Baiting technique. Field samples of submerged substrates (plant matter, sediment, debris), surface scum,
and source water were collected in sterile Whirl-Pak® bags. Samples were refrigerated if not treated the
same day. Within three days, collected material was transferred by sterile tweezers and/or pipette to a 6 cm
diameter Petri dish or a 6 cm square dish containing a few ml of bottled FIJI® Water (Fig. 1). Next, several
thousand Lecane inermis in 3 mL of rotifer culture and a floating plastic coverslip (Fisherbrand™
disposable cover slips, 22 × 22 mm) as potential substrate for Sommerstorffia, were added to each dish.
Coverslips were monitored in situ for up to seven days with a compound microscope (Fig. 2).
The bait rotifer Lecane inermis was isolated in 2013 from a spring in Florence, Alabama. Derived
from a clone of a single individual washed through several drops of bottled spring water, Lecane inermis
was cultured in 10 cm diameter Petri dishes containing 25 ml of FIJI Water and three rice grains (Mahatma
basmati rice) or 0.25 ml of V-8 Juice. Rotifers consumed unidentified bacteria and particulate debris in the
cultures. Subcultures were made at least every three weeks by inoculating sterile dishes with 3 ml of an
existing culture. Segars (1994-95) was used to identify the rotifer species.
Producing single spore isolates. Coverslips bearing Sommerstorffia were placed specimen side up on a
glass pedestal (Figs. 3-4), and several drops of rotifer culture were added forming a modified sitting drop
culture that is here called a standing pool culture. After three days in a moist chamber the standing pool
was rinsed with distilled water in an attempt to eliminate infusoria and a few drops of rotifer culture were
added. After 8-12 hours the standing pool was rinsed again to remove uncaptured rotifers. The standing
pool was monitored through a compound microscope over several days for the development of mature
sporelings in the surface film. It was sometimes necessary to repeat the process of adding rotifers followed
by rinsing to acquire numerous sporelings in the pool’s surface. A few drops of rotifer culture were then
added and the surface immediately monitoring through a dissecting microscope for sporeling-trapped
Phytologia (Dec 20, 2024) 106(4) 53
rotifers (detectable by the rotifer’s body writhing while the rotifer’s head remained stationary). Within 15
minutes of being trapped, rotifers were picked up with a flame sterilized Irwin loop (Schram and Davison
2012) and washed through several separate drops of FIJI Water. The final transfer was to a sitting drop of
FIJI Water (Fig. 5) to which several drops of rotifer culture were then added. This sitting drop culture was
held in a moist chamber and monitored for up to five days. Sitting drop cultures with mature hyphae and/or
sporelings were transferred as floating coverslips to individual 10 cm diameter Petri dishes freshly prepared
with three rice grains, 3 ml of rotifer culture, and 25 ml of FIJI Water.
Maintenance of Sommerstorffia cultures. Cultures derived from single spore isolates were kept at room
conditions and subcultured every 30 days as follows. Sommerstorffia-infected plastic coverslips (or
fragments of plastic coverslips) bearing hyphal pegs and/or sporelings were transferred to freshly prepared
Petri dishes containing three rice grains, 3 ml of rotifer culture, and 25 ml of FIJI Water.
Photography and Measurements. Micrographs were taken through a Nikon Alphaphot-2 YS2 or an
Eclipse E200 compound microscope, fitted with a Handycam HDV 1080i camera with an adaptor from
Martin Microscope Company or an iPhone 14 Pro with LabCam® adaptor from iDu Optics®.
Measurements were made from water-mounted Sommerstorffia in the living state. Imaged material was
attached to plastic or glass coverslips floated in Petri dish cultures (slide mounted for photomicroscopy).
Glass coverslips were held in the surface water of cultures by a plastic bracket that prevented the glass from
sinking (Fig. 6). Measurements were made from digital photographs in ImageJ (https://imagej.net/ij/)
calibrated by images of a stage micrometer taken at same scale. Distracting background artifacts were
removed with Adobe Photoshop in some images.
Preservation of dried voucher specimens. Plastic coverslips bearing Sommerstorffia in various stages of
development were dried specimen side up on filter paper in a vented Petri dish. Following Wu et al. (2004),
Petri dishes were placed in archival quality paper packets with attached labels. Specimens were deposited
in the Kriebel Herbarium (PUL) at Purdue University, the Yale University Herbarium (YU), and the
University of North Alabama Herbarium (UNAF).
Molecular Methods. Samples for DNA extraction were processed from Petri dish cultures, which were
decanted by pipetting, leaving surface sporelings and thalli in the dish with minimal water. A sterile razor
blade was used to scrape the dish and concentrate material to one side. Approximately 1 ml volume was
pipetted into a 1.7 ml sterile tube and centrifuged at 10,000 rpm for 3 min, after which a pellet was visible
in the bottom of the tube. The supernatant was removed, and the tube with pellet was placed in an ultralow
freezer at -80°C. After 5 min, the frozen pellet was ground by hand using a disposable plastic pestle to
mechanically disrupt the cells. The 1.7 ml tube was then utilized for the first steps of DNA extraction using
a Qiagen DNeasy® Plant Mini Kit (catalog no. 69104), following the Quick Start Protocol (QIAGEN).
PCR reactions used a total volume of 50 uL including 4 uL template DNA, 25uL Promega GoTaq®
Green Master Mix (catalog no. M7122), 2.5uL each primer (0.3uM final concentration), and 16 uL
nuclease-free water. For COI, primer pairs were OomCoxI-Levup – OomCoxI-Levlo and OomCoxI-Levup
– FM85mod (Robideau et al. 2011). For ITS, primer pairs were ITS1oo–ITS4(r) and ITS3oo–ITS4(r)
(White et al. 1990; Ritt et al. 2016, 2018). A Techne 3Prime G Thermo Cycler (Bibby Scientific) was used
for the following conditions: (1) an initial denaturation of 60 s at 94°C, (2) 35 cycles of 60 s at 95°C, 30 s
at 55°C, 60 s at 72°C, and (3) a final extension of 60 s at 72°C. PCR products were verified for
appropriately-sized bands on a 2% agarose gel stained with EtBr; electrophoresis was for 40 min at 70 W.
Products were shipped to Eurofins Genomics, Louisville, KY, USA for PCR purification and Sanger
sequencing of both forward and reverse strands using the identical forward and reverse PCR primers listed
above.
Specimens with data for both COI and ITS markers from Verrucalvaceae and Saprolegniaceae,
along with outgroup taxa Apodachlya brachynema and A. minima (Robideau et al. 2011; Rocha et al. 2018),
Phytologia (Dec 20, 2024) 106(4)
54
were downloaded from GenBank. Sequencing results were edited and aligned with GenBank taxa in
Geneious Prime 2023.2.1 producing an alignment of 19 sequences with 1,583 characters (680 for COI and
903 for ITS). New Sommerstorffia DNA sequences were deposited in GenBank, with accession numbers
PP404027–PP404033 for COI and PP407290–PP407296 for ITS (Table 1). The software ModelFinder
(Kalyaanamoorthy et al. 2017) was used to determine the optimum model of nucleotide substitution for
each genetic marker and performed a maximum likelihood (ML) analysis on the partitioned dataset in IQ-
TREE (Trifinopoulos et al. 2016). Settings for the best-fit substitution model were TPM2+F+G4 (COI) and
TVM+F+I+G4 (ITS), with ultrafast bootstrap support analysis using 1000 alignments (Hoang et al. 2018).
The resulting ML tree was modified for viewing with iTOL (Letunic and Bork 2021).
Table 1. Taxa included in phylogenetic analyses including reference/locality, isolate, and GenBank
accession numbers for specimens. Data downloaded from GenBank includes taxa from Robideau et al.
(2011). Taxa and isolates correspond to specimens in Figure 7.
Oomycete Taxa
Reference / Locality
(State: County)
Isolate
GenBank Accession
Number
COI
ITS
Apodachlya brachynema
Robideau et al. 2011
CBS 184.82
HQ708197
HQ643124
Apodachlya minima
Robideau et al. 2011
CBS 185.82
HQ708199
HQ643126
Pythiopsis terrestris
Robideau et al. 2011
CBS 110059
HQ708453
HQ643406
Achlya papillosa
Robideau et al. 2011
CBS 101.52
HQ708175
HQ643102
Achlya spinosa
Robideau et al. 2011
CBS 576.67
HQ708182
HQ643109
Leptolegnia caudata
Robideau et al. 2011
CBS 680.69
HQ708211
HQ643137
Protoachlya paradoxa
Robideau et al. 2011
CBS 100.44
HQ708451
HQ643404
Saprolegnia ferax
Robideau et al. 2011
BR 114
HQ709029
HQ643988
Saprolegnia parasitica
Robideau et al. 2011
CBS 113187
HQ709046
HQ644005
Plectospira myriandra
Robideau et al. 2011
CBS 523.87
HQ708449
HQ643402
Aphanomyces cochlioides
Robideau et al. 2011
CBS 477.71
HQ708188
HQ643115
Aphanomyces cladogamus
Robideau et al. 2011
CBS 108.29
HQ708186
HQ643113
Sommerstorffia pugioniformis sp.
nov.
CT: New Haven
CT03
PP404028
PP407291
Sommerstorffia pugioniformis sp.
nov.
CT: New Haven
CT04
PP404029
PP407292
Sommerstorffia spinosa
CT: New Haven
CT01
PP404027
PP407290
Sommerstorffia spinosa
FL: Taylor
SP05
PP404030
PP407293
Sommerstorffia spinosa
FL: Sarasota
SP06
PP404031
PP407294
Sommerstorffia spinosa
AL: Lawrence
SP07
PP404032
PP407295
Sommerstorffia spinosa
AL: Lauderdale
SP08
PP404033
PP407296
Phytologia (Dec 20, 2024) 106(4) 55
RESULTS
Phylogenetic analysis. The final alignment contained 464 parsimony-informative characters (COI: 122
characters, ITS: 342 characters). Maximum Likelihood analysis recovered all Sommerstorffia samples in a
well-supported clade (100% bootstrap value), sister to two other genera within Verrucalvaceae, and more
distantly related to species recognized as members of Saprolegniaceae (Fig. 7). Sommerstorffia sp. nov. in
Connecticut, USA was recovered as a well-supported clade divergent from the sympatric (and syntopic) S.
spinosa in Connecticut. The widely disjunct (geographic distances of ca. 1,700-2,000 km) samples of S.
spinosa from Connecticut, Alabama, and Florida formed a well-supported clade. Uncorrected sequence
divergence between Sommerstorffia species was 3.5–2.7%, with intraspecific divergences of 0.4% (S.
pugioniformis sp. nov.) and 0.4–1.6% (S. spinosa).
TAXONOMY
Sommerstorffia pugioniformis P.G. Davison, J.M. Ray, and C.A. Davison, sp. nov. Figs. 8 – 21, 27.
MycoBank no.: MB 855653
Typification: USA, Connecticut, New Haven County, New Haven, East Rock Park, 41.33292°N,
72.90932°W, 0.6 m, from a sample of water with submerged vegetation and sediment collected
along margin of Mill River by P.G. and C.A. Davison, June 6, 2020; strain CT03 isolated by P.G.
Davison. HOLOTYPE Fig. 8, this publication.
Etymology: The specific epithet refers to the sporeling being dagger like; from the Latin words pugio =
dagger and forma = shape.
Diagnosis: Similar to Sommerstorffia spinosa, however, Sommerstorffia pugioniformis differs from S.
spinosa as follows: the neck of the sporeling is 1.2–2 times longer than the length of the base of the
sporeling (versus the neck about half the length of the base); the neck of the sporeling tapers ending in a
minute subglobose apex (versus neck not tapered, ending in a blunt tip). Additionally, the hyphal pegs of
S. pugioniformis are more narrowly pointed, being 1 µm wide (versus 2–5 µm wide).
Description: Thallus external to host rotifer consisting of tubular hyphae, (1–)2–5(–6) branched, branches
becoming more numerous with subsequent rotifer captures; hyphal branches 35–76 µm long × 3.3–5.0 µm
wide; base of branches sometimes contorted in zig-zag fashion, region where two or more branches
originate sometimes swollen to 7–14 µm wide; distal ends of branches sweeping away from substrate;
branch tips ending in a predaceous peg 3.0–5.2 µm long × 0.9–1.2 µm wide, often with a thin extrusion of
unknown material extending 2–4 µm beyond the apex, pegs sometime slightly swollen at the apex. Thallus
inside host rotifer a lobed zoosporangium filling wholly, or in part, the rotifer body, the lobes up to 17 µm
in diameter. Exit tubes 14–70 µm long × 4–6 µm wide. Cystospores 6–9 µm diameter, numbering 12–60
atop the exit tube. Zoospores laterally biflagellate, reniform, 6.0–8.6 × 5.2–6.8 µm (measured in rocking
phase atop exit tube). Sporelings consisting of a swollen base and a neck that serves as a predaceous peg;
the base lemon-shaped to subglobose, 6.8–10 µm long × 5.1–7.5 µm wide, often with a narrow, transverse,
irregular band of clear cytoplasm; the neck 11.1–17.3(–21.7) µm long × 2.0–2.7 µm wide, tapering in the
distal ¼–½ to a short, narrow isthmus 0.2–0.6 µm wide tipped by a subglobose head 0.7–1.1 µm in diameter.
Zoospores encysting twice, once atop the exit tube of zoosporangium and again after flagellated dispersal;
second cyst attached to submerged surfaces or to the thin film at the water’s surface. Second cyst releases
a sporeling that remains attached to substrate. Oogonia terminating 12–74 µm long hyphal branches,
bullate, 23–30 µm outer diameter. Oospheres single, usually not maturing, 17–20 µm diameter. Antheridia
not seen.
Phytologia (Dec 20, 2024) 106(4)
56
Geographical distribution: North America, USA, Connecticut, New Haven.
Habitat: Known only from shallow, bankside water in the Mill River downstream of the Lake Whitney
dam. See Fig. 29 for photograph of the type locality.
Host selection: cf. Lepadella, Lecane inermis.
Remarks: Development of oogonia was sporadic and restricted to older cultures. Almost all development
can be classified as endo-epibiotic as described by Karling (1952) in which both an internal zoosporangium
and external hyphae bearing pegs form from a sporeling that has captured a rotifer. Rarely, as noted by
Karling for S. spinosa, development is completely endobiotic (internal zoosporangium but no external
hyphae) (Fig. 18) or epibiotic (external hyphae form but no zoosporangium).
As in S. spinosa, there is often an extension of hyaline material (Fig. 10) that protrudes beyond the
predaceous pegs. Arnaudov and Damova (1947-1948) first noticed these extensions and questioned their
purpose. Karling (1952) stated the material was not adhesive to passing rotifers and the purpose of the
extensions was unknown. Prowse (1954), unconvincingly, concluded the material was mucilage that
appeared to capture rotifers. Having observed the grazing habits of Lecane inermis actively chewing and
completely consuming in cannibalistic fashion the ghostly remains of partially decomposed rotifers and
membranous material in V-8 juice, a plausible explanation for these extensions is that they serve to
stimulate the grazing activity of the rotifer leading it to the predaceous peg. Only by biting the peg is the
rotifer trapped. We hypothesize that the hyaline extensions are an adaptation that improves the efficacy of
the hyphal pegs in trapping rotifers.
Additional specimens examined: USA. Connecticut: New Haven County, New Haven, Mill River, East
Rock Park, 41.33292°N, 72.90932°W, 0.6 m, submerged vegetation and sediment, 7 Jun 2021, P.G. and
C.A. Davison, occurring sympatrically with S. spinosa, no specimen preserved; same data except
41.32614°N, W 72.90701°W, 0.6 m, patch of surface scum (few cm in diameter) 0.5 m from shore, preying
on native cf. Lepadella, 7 Jun 2021, P.G. and C.A. Davison 2021-06-07.2 (PUL, UNAF, YU)
Sommerstorffia spinosa Arnautov, Flora (Regensburg) 116: 109 (1923a), Figs. 22 – 26, 28
Lectotype designated by Dick (2001), Flora, Jena 116: 109-113, figs 1-5 (Arnaudov 1923a)
Description: see Karling (1952, 1966).
Geographical distribution: ASIA. Japan (Saikawa and Hoshino 1986; Saikawa et al. 2021), India (Karling
1966) AUSTRALASIA. New Zealand (Karling 1968), EUROPE. Bulgaria (Arnaudov 1923a,b),
Czechoslovakia (Cejp 1959 fide citation in Johnson et al. 2002); England (Prowse 1954), Poland (Próba
1980, Czeczuga and Próba 1980; Czeczuga et al. 2015 ); Russia (https://www.inaturalist.org/
observations/163541892). NORTH AMERICA. USA. Alaska, Louisiana, New Jersey (Karling 1952),
Virginia (Karling 1952; Miller 1965), Massachusetts (Sparrow 1929), New York (Sparrow 1933; this
study), Alabama, Florida, Mississippi, North Carolina, Tennessee (this study), West Indies, Cuba (Sparrow
1952).
Habitat: Freshwater ponds, lakes, springs, rivers, soil (Arnaudov 1923a; Czeczuga et al. 2000; Karling
1952, 1966, 1968; Próba 1980; Saikawa and Hoshino 1986). See Fig. 30 for photograph of the type locality.
Native substrates as reported in the literature (substrates used as bait not included): Cladophora (Arnaudov
1923a, Prowse 1954, Sparrow 1933), Rhizoclonium (Sparrow 1929), littoral detritus (Saikawa and Hoshino
1986), soil—presumably aquatic sediment—(Karling 1952, 1966, 1968). Native substrates with attached
thalli observed in baited dishes (this study): Ceratophyllum demersum, Chara sp., Fissidens fontanus, Najas
guadalupensis, Podostemum ceratophyllum, Spirodela polyrhiza, Utricularia sp.
Phytologia (Dec 20, 2024) 106(4) 57
Host selection: Rotifers: Lecane sp., Lecane inermis (this study), Colurella sp. and Lepadella sp. (Glocking
et al. 2014); protists: Entosiphon ovatum (Karling 1952), Difflugia pyriformis (Saikawa et al. 2021).
Remarks: So far, oogonia in S. spinosa have only been seen in European material (Arnautov 1923a;
Czeczuga and Próba 1980; Próba 1980; Czeczuga et al. 2000). Karling (1952) did not observe oogonia in
the Alaskan material he maintained in culture for over a year. Likewise, oogonia have not been seen in any
of our isolates from seven U.S. states maintained in culture for 1–4 years.
Additional specimens examined: USA. Alabama: Colbert County, Cane Creek boat ramp, 34.75422°N,
87.86181°W, 126 m, Spirodela polyrhiza floating between cattail stalks, 7 Aug 2020, P.G. Davison, no
specimen preserved; Tennessee River just upstream of confluence with Cane Creek, 34.75554°N,
87.86135°W, 126 m, Najas guadalupensis submerged near shoreline, 7 Aug 2020, P.G. Davison, no
specimen preserved; Sheffield, Tennessee River, 34.76739°N, 87.71145°W, 126 m, algae attached to
underpinning of floating dock, 12 Aug 2020, P.G. Davison, no specimen preserved; Lauderdale County,
Florence, King Spring, 34.85683°N, 87.65412°W, 165 m, submerged dialysis tubing, Feb 2011, P.G.
Davison, no specimen preserved; Big Spring north of Hwy 20, 34.78945°N, 87.68301°W, 128 m, duckweed
community baited with poppy seeds, Jan 2013, P.G. Davison, preying on Lecane inermis, no specimen
preserved; unnamed stream on the campus of the University of North Alabama, submerged Fissidens
fontanus, 34.80643°N, 87.68607°W, 150 m, 24 Dec 2020, P.G. Davison 2020-12-24.1 (PUL, UNAF, YU);
Wildwood Park, Cypress Creek, 34.80370°N, 87.69524°W, 128 m, lower stems of Podostemum
ceratophyllum, 3 Nov 2023, P.G. Davison 2023-11-03.1 (PUL, UNAF, YU); Wildwood Park, upland
headwater spring under forest cover, 34.81113°N, 87.69456°W, 150 m, submerged brown detritus, 8 Jul
2023, P.G. Davison, no specimen preserved; McFarland Park, floodplain of Tennessee River, golf course
pond, 34.78670°N, 87.68186°W, 127 m, Spirodela polyrhiza floating on pond, 5 Jul 2023, P.G. Davison,
no specimen preserved; McFarland Park, slough along Hwy 20, 34.78805°N, 87.68555°W, 127 m, Lemna
sp. and Landoltia punctata floating in surface, 5 Jul 2023, P.G. Davison, no specimen preserved; Lawrence
County, Prairie Grove Glade Preserve, 34.51724°N, 87.50322°W, 180 m, Chara in small pond, 24 Mar
2021, P.G. Davison 2021-03-24.1 (PUL, UNAF, YU); Connecticut: New Haven County, New Haven,
Mill River, East Rock Park, 41.33292°N, 72.90932°W, 0.6 m, submerged vegetation and sediment, 7 June
2021, P.G. and C.A. Davison 2021-06-07.1 (PUL, UNAF, YU); Regional Water Authority Recreational
Area, 41.33579°N, 72.91281°W, 1 m, Spirodela polyrhiza floating on pond, 6 Jun 2020, P.G. and C.A.
Davison, no specimen preserved; Florida: Taylor County, Steinhatchee River, 29.67580°N, 83.36589°W,
0.5 m, floating and submerged woody debris, 29 Jul 2021, P.G. Davison 2021-07-29.1 (PUL, UNAF, YU);
Sarasota County, Lake Jervey, State College of Florida Venice campus, 27.04335°N, 82.31958°W, 1 m,
submerged debris, 26 Dec 2020, E.L. Kasl 2020-12-26.1 (PUL, UNAF, YU); Sarasota Golf Community,
roadside ponds, 27.03914°N, 82.38327°W, and 27.03635°N, 82.36362°W, 4 m, submerged debris, 26 Dec
2020, E.L. Kasl, no specimens preserved; Mississippi: Tishomingo County, Tishomingo State Park,
Haynes Lake, 34.61559°N, 88.19349°W, 145 m, Chara and sediment, 17 Jul 2023, P.G. Davison 2023-07-
17.1 (PUL, UNAF, YU); New York: Putnam County, Town of Kent, Veterans Memorial Park,
41.44194°N, 73.70960°W, 177 m, Ceratophyllum, Potamogeton, and debris, 11 Jun 2022, P.G. Davison
2022-06-11.1 (PUL, UNAF, YU). North Carolina: Gaston County, Crowders Mountain State Park, Shorts
Lake, N 35.20761°N, W 81.29243°W, 252 m, Utricularia and sediment, 16 Mar 2024, P.G. Davison 2024-
03-16.1 (PUL, UNAF, YU); Watauga County, Blowing Rock, Mayview Lake, 36.13370°N, 81.68141°W,
1074 m, surface scum and submerged Callitriche, 28 Jul 2022, P.G. Davison 2022-07-28.1 (PUL, UNAF,
YU); Tennessee: Hawkins County, Holston River at Surgoinsville, 36.46853°N, 82.85095°W, 333 m,
submerged leaves of Vallisneria americana, 18 Oct 2023, D.H. Webb, no specimen preserved; Lawrence
County: Holly Creek near confluence with Shoal Creek, 35.02527°N, 87.57945°W, 165 m, exposed,
submerged roots of Justicia americana, 1 Aug 2023, P.G. Davison and J.M. Ray 2023-08-01.1 (PUL,
UNAF, YU).
Phytologia (Dec 20, 2024) 106(4)
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DISCUSSION
Based on both morphological and phylogenetic analyses, S. pugioniformis is described as distinct
species within a previously monotypic genus. The phylogenetic position of Sommerstorffia within the
Verrucalvaceae (Beakes and Thines 2017; Rocha et al. 2018) is confirmed using data from the two
recommended genetic markers (Robideau et al. 2011).
When detectable, Sommerstorffia was found attached to floating coverslips 3 to 6 days after the
addition of bait rotifers. As early as one day after baiting, rotifers may be seen head planted on the surface
of opaque, submerged material where it is difficult to see evidence of a specific predator. The rotifer’s
body projecting away from the substrate may be seen in profile. Such instances are often indicative of the
rotifer having been trapped by either Sommerstorffia or the zygomycete fungus Zoophagus. From our
observations and others (Arnaudov 1923b; Sparrow 1933; Saikawa et al. 2021) Sommerstorffia spinosa and
Zoophagus cf. insidians frequently co-occur. Either predator may appear on the dish bottom or attached to
submerged plants or debris. Only Sommerstorrfia readily spreads by zoospores and soon appears with
trapped rotifers under the transparent coverslip where its distinctive morphology is readily observed.
As a precedent to the utility of the bait rotifer, L. inermis is widely adopted by others, notably in
studies of rotifer-fungal interactions (Fiałkowska and Pajdak-Stós 2018; Fiałkowska et al. 2020) and for
the culture of carnivorous tardigrades (Roszkowska et al. 2021; Tůmová et al. 2022). Tardigradologists
isolated L. inermis from the wild (Suzuki 2003) or obtained a culture from another lab (Tůmová et al. 2022).
Lecane inermis reproduces primarily by parthenogenesis and is known to have a high reproductive capacity
(Edmondson 1946; Kowalska et al. 2017). Other rotifer species with the grazing habit of feeding may be
suitable to culture as bait; however, we have come to rely on L. inermis for its ease of culturing and high
fecundity. Using a Sedgewick-Rafter counting cell, we estimated from a seven-day old, V-8 juice, L.
inermis culture that the 3 ml added to field samples or culture dishes supplies between 6,350 and 11,000
rotifers, not counting rotifer eggs.
We speculate, on the basis of the narrow hyphal pegs, that Sparrow (1929) in his Figure 1 of S.
spinosa may have illustrated S. pugioniformis. This specimen was collected from “a small pond in Fresh
Pond Parkway, Cambridge, Mass.” located ca. 6 km NW of the Atlantic coastal city of Boston. We
collected S. pugioniformis from a freshwater stream 4 km inland from the Atlantic coast in Connecticut.
Our collecting efforts and the highly speculative reidentification of Sparrow’s Massachusetts specimen
suggests the new species may be restricted to near estuarine habitats along the Atlantic coast; however, the
targeted collecting effort to date is inadequate to determine with any confidence the geographic and
physiographic distribution of the new species.
ACKNOWLEDGEMENTS
We thank Teodor T. Denchev and Cvetomir M. Denchev for sending publications by Arnaudov,
Teodor T. Denchev for sending the photo of Dragichevsko Blato Swamp, Peter Döbbeler for formulating
the Latin binomial Sommerstorffia pugioniformis, Sally Glockling for sharing methods of
micromanipulations, Emily Kasl for samples from Florida, the late David H. Webb for a sample from
Tennessee, and the following reviewers for their careful attention to the manuscript: William J. Davis, Alvin
Diamond, and D. Rabern Simmons. The University of North Alabama Department of Biology provided
financial support for the molecular laboratory portion of this research.
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Figures 1-6. Materials and methods. 1, arrangement of opaque material in baited dish; 2, in situ observation
of floating coverslip in surface scum including duckweeds; 3-4, mount of standing pool culture—the glass
pedestal prevents water at the edge of the coverslip from being pulled under the coverslip by capillary
forces; 5, sitting drop culture on plastic coverslip; 6, glass coverslip held in surface water by a submerged
plastic bracket (folded fragment of a Rinzl plastic slide). With proper adjustment of the water’s depth, the
adhesive force of water pulls the coverslip firmly against the rails of the bracket and adheres the bracket to
the dish bottom.
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Figure 7. Maximum Likelihood phylogeny of Oomycote species from GenBank (Robideau et al. 2011) and
Sommerstorffia isolates generated in this study. Analysis based on a partitioned data set of DNA sequences
for COI and ITS with bootstrap support values of >90% shown on basal nodes. Specimens correspond to
taxa and isolates in Table 1.
Phytologia (Dec 20, 2024) 106(4) 63
Figures 8-14. Sommerstorffia pugioniformis. 8, 10, 12, 13, strain CT03; 9, 11, 14, strain CT04. 8, external
thallus developed from a sporeling that captured the rotifer at left, consisting of four hyphal branches, one
branch with a captured rotifer; mature sporelings at arrows; 9, a three-branched thallus developed from a
sporeling-captured rotifer; 10, hyphal peg with extension of hyaline material at arrow; 11, thallus with
multiple branches and ghostly remains of several trapped rotifers; 12, terminal pegs at end of branches that
sweep away from substrate; 13, three-branched external thallus developed from basal part of sporeling at
arrowhead, the neck of sporeling (at arrow) ends in the jaws of rotifer; 14 three-branched external thallus
developed from a sporeling-captured rotifer, base of capturing sporeling at arrowhead. Scale bar = 50 µm
8, 9, 11, 12, 14; 10 µm 10, 13.
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Figures 15-21. Sommerstorffia pugioniformis. 16, 17, 19, 21, strain CT03; 15, 18, 20, CT04. 15, mature
sporeling with swollen base and slender neck, a rotifer bites the tapered end of the neck that bears a minute,
swollen tip and becomes trapped; 16, three sporelings in two focal planes, at left focused on clear band in
cytoplasm of the swollen base, at right focused on the necks of the sporelings and empty cyst walls at
arrowheads; 17, mature sporelings with adjacent hyaline cyst walls, necks bent where joining the base by
coverslip pressure; 18, endobiotic development lacking external thallus except for the exit tube leading
from a lobe of the vacated zoosporangium, zoospores at arrowhead actively departing from the cystospore
cluster atop the exit tube; 19, lobed zoosporangium yet to form zoospores, formation of exit tube at
arrowhead; 20, oogonium on long hypha of older thallus; 21, two oogonia on shorter hyphae hidden from
view. Scale bar = 10 µm 15, 16, 17, 19; 50 µm 18, 20, 21.
Phytologia (Dec 20, 2024) 106(4) 65
Figures 22-26. Sommerstorffia spinosa. Strain CT01. 22, external thallus of two branches ending in
mature pegs with curled hyaline extensions, thallus developed from a sporeling that captured the rotifer
now partially decomposed, cluster of cystospores atop exit tube from posterior end of rotifer; 23, external
thallus of three branches ending in mature pegs, thallus developed from sporeling-captured rotifer whose
body is filled with a vacated zoosporangium, exit tube (arrowhead) departing zoosporangium at anterior
end of rotifer; 24, mature peg at tip of hypha bearing hyaline extension at arrow; 25-26, mature sporelings
with short, nontapered necks, cyst wall at arrows. Scale bar = 50 µm 22-23, 10 µm 24-26.
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Figures 27-28. Sommerstorffia developing under glass coverslips. 27, S. pugioniformis strain CT04; 28 S.
spinosa strain CT01. Scale bar = 50 µm.
Phytologia (Dec 20, 2024) 106(4) 67
Figures 29-30. Type localities. 29, type locality of Sommerstorffia pugioniformis, Mill River, Connecticut,
June 6, 2020; 30, type locality of Sommerstorffia spinosa, Dragichevsko Blato Swamp in Mt. Lyulin,
Bulgaria, May 9, 2014 (photo taken by Teodor T. Denchev).
