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Chapter 13
Phylum Rotifera
Robert L. Wallace
Department of Biology, Ripon College, Ripon, Wisconsin, USA
Terry W. Snell
School of Biology, Georgia Institute of Technology, Atlanta, Georgia, USA
Hilary A. Smith
Department of Biological Sciences, University of Notre Dame, Notre Dame, Indiana, USA
Chapter Outline
Introduction to Rotifera
225
General Characteristics
225
Biogeography228
Evolutionary Relationships
230
General Biology
231
External Morphology
231
Organ System Structure and Function
232
Corona232
Trophi and Gut
232
Muscular System
235
Neural System
236
Excretory System: Protonephridium
236
Reproductive System
237
Environmental Physiology
238
Locomotion238
Physiological Ecology
238
Environmental Toxicology
240
Anhydrobiosis240
Generalized Stress Responses
242
Ecology and Evolution
242
Diversity and Distribution
242
Phenotypic Variation
242
INTRODUCTION TO ROTIFERA
General Characteristics
Phylum Rotifera comprises approximately 2000 species of
unsegmented, bilaterally symmetrical invertebrates, most of
which are found in freshwaters (Clément and Wurdak, 1991;
Distribution and Population Movements
243
Colonial Rotifers
244
Sessile Rotifers
246
Reproduction and Life History
249
Reproduction249
Aging252
Population Dynamics
253
Ecological Interactions
258
Foraging Behavior
258
Functional Role in the Ecosystem
259
Competition with Other Zooplankton
261
Predator–Prey Interactions
261
Parasitism on Rotifers
264
Rotifers as Parasites
265
Collecting, Culturing, and Preparation for Identification
265
Collecting265
Culturing266
Laboratory Culture
266
Aquaculture267
Preparation for Identification
268
References269
Wallace et al., 2006; Segers, 2007). Their size ranges from 40
to 2000 μm, the smallest being only about 6 times the diameter of a human red blood cell. Because of their size, shape,
and habitat, rotifers can be confused with protozoans (protists) (Chapter 7) and gastrotrichs (Chapter 12), but those taxa
do not possess jaws and their ciliation is not distributed in the
Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00013-9
Copyright © 2015 Elsevier Inc. All rights reserved.
225
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same way as in rotifers. Very few rotifers are parasitic; nearly
all are either raptorial predators or microphagous suspension feeders or grazers. Collectively this phylum is widely
dispersed, being found in all types of freshwater habitats at
densities up to about 1000 individuals per liter. With sufficient food, populations may surpass 5000 individuals per liter
(Feike and Heerkloss, 2009). In some rather unusual water
bodies, exceedingly large populations can develop; sewage
ponds may contain about 12,000 per liter (Seaman et al.,
1986), and soda water bodies in Chad can hold well over
100,000 per liter (Iltis and Riou-Duvat, 1971). Similar populations can be obtained in small chemostat systems (Boraas,
1983), but aquaculture systems population levels above 107
individuals per liter have been reported (Park et al., 2001).
These tiny animals possess two conspicuous features.
First, a specialized ciliated region called the corona (L.,
crown) caps the anterior end. The corona is commonly composed of two concentric rings of cilia (Figures 13.1–13.2).
When viewing the corona of many species, one often is
struck with the impression of a rotating wheel. This image
comes from the metachronal (rhythmic and sequential) beating of their cilia, and inspired early microscopists with the
name for the phylum (L., rota, wheel and L., ferre, to bear):
the wheel-bearers. In free-swimming species the corona is
used in locomotion, but all species employ it in some way
to collect food. However, in adults of some species ciliation
is lacking and the corona is funnel or bowl-shaped, with the
Protozoa to Tardigrada
mouth located at the bottom. The second obvious feature that
all rotifers possess is a muscular pharynx, termed the mastax,
that includes a complex set of jaws called trophi (G., troph,
nourish). In some rotifers, the trophi are so unique that taxonomists distinguish species by critical morphological features
of these minute structures (see the section “Trophi and Gut”).
Although most rotifers inhabit freshwaters, some genera
also have members that occur in saline waters. For example,
21 of the 39 species in the genus Synchaeta are known to
occur in brackish to full-strength marine waters (Segers,
2007). However, only about 100 species distributed among
22 genera in the phylum are found exclusively in marine
habitats (Ricci and Fontaneto, 2003). In general, rotifers
are not as diverse or as abundant in marine environments
as microcrustaceans, but they occur in estuarine waters, as
well as in interstitial, tide pool, and near-shore marine habitats. Occasionally rotifers comprise an important portion of
the biomass of marine zooplankton (Dolan and Gallegos,
1992). Inland saline waters, termed athalassohaline, are also
habitats for rotifers (Segers, 2007; Walsh et al., 2008).
Remarkably, some rotifers are found at the interface bridging aquatic and terrestrial habitats, i.e., they inhabit the film
of water covering mosses, lichens, and liverworts. This habitat, referred to as limnoterrestrial (Figure 13.3(a)–(b)), is also
home to nematodes (Chapter 14) and tardigrades (Chapter 17).
Additionally, rotifers are members of pitcher plant and treehole
communities, the phytotelmata (Figure 13.3(c)–(d)), and can be
FIGURE 13.1 Lateral view of a generalized rotifer. Modified with permission from Koste and Shiel (1987).
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Phylum Rotifera
found in containers holding water, such as birdbaths, as well as
in discarded cups and tires stored outdoors (Figure 13.3(e)–(g)).
Furthermore, rotifers often are abundant in the interstitial water
of soils and sediments (Pourriot, 1979) including peat (Błędzki
and Ellison, 2002). Depending on the soil type and its moisture
level estimates of their densities range from about 32,000 to
more than 2 million per square meter. Because of their high
population levels and rapid metabolism, rotifers probably play
an important role in nutrient cycling in soils.
In addition to the variation in their habitats, the diversity of rotifer life histories is remarkable. Most are motile,
either swimming as members of the plankton or crawling
over plants or within sediments. However, after a brief,
free-swimming stage, juveniles in three families of sessile
rotifers attach permanently to a substrate, usually a freshwater plant (Wallace, 1980). The vast majority of rotifers
are solitary, but about 25 species form colonies of various
sizes (Wallace, 1987). Higher taxonomic groups are largely
known for their differences in reproductive strategies.
Two classes of rotifers are recognized: class Pararotatoria, comprising a single small family Seisonidae; and class
Eurotatoria, containing subclasses Bdelloidea and Monogononta (Segers, 2002; Wallace et al., 2006). Seisonids
are exclusively marine and obligatorily sexual. They are
not discussed in detail here. All bdelloids are exclusively
parthenogenetic, whereas monogononts are intermittently
sexual; that is, they are cyclical parthenogens (see “Reproduction and Life History”). However, a complication to
these generalizations is that males have never been reported
for some monogononts. The variety of form (Figures 13.4
and 13.5) and life histories within the phylum offers a rich
field of study.
Additional accounts of this phylum may be found in
most texts of general and invertebrate zoology, in some
specialized books about inland waters (Wallace and Ricci,
2002; Wallace and Smith, 2009), or in advanced texts
(Edmondson, 1959; Ruttner-Kolisko, 1974; Wallace et al.,
2006). In the 1800s, some beautifully illustrated works were
published that still offer an excellent depiction of these animals, although the taxonomy of some species is out of date
(Hudson and Gosse, 1886). The digital images provided by
Jersabek et al. (2003) of permanent slides made by Frank
J. Myers are instructive for the diversity of animals covered, as well as for their historic value. There is no single
scientific journal or set of journals in which researchers
publish their research on rotifers; the field simply is too
FIGURE 13.2 Female and male Brachionus species. Modified with permission from Pourriot and Francez (1986).
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228
(a)
(b)
(c)
(d)
(e)
(f)
Protozoa to Tardigrada
(g)
FIGURE 13.3 Little-known habitats for rotifers. Two limnoterrestrial habitats: (a) Sphagnum moss in a bog pond; (b) lichen on a granite outcrop. Two
phytotelma: (c) Sarracenia purpurea, the northern pitcher plant); (d) treeholes. Three container habitats: (e) birdbath; (f) discarded cup; (g) discarded tires.
((a)–(c), (e)–(g), R.L. Wallace; (d), courtesy of Christian Jersabek, University of Salzburg.
diverse. However, since 1976, a small group of researchers
(ca. 35–135) has gathered every 3 years to hold the International Rotifer Symposium. (At the time of this writing
13 such meetings have been held and their symposia volumes published.) Some of the papers discussed in this chapter were presented at the rotifer meetings. Several Internet
sites describe rotifers and post stunning photomicrographs.
However, we urge care in using keys posted on the Internet, as they are commonly based on regional samples, and
identification of specimens based on photographs or line
drawings alone is unwise. A detailed coverage of the phylum and of specific taxonomic groups is available in the
Series Guides to the Identification of Microinvertebrates of
the Continental Waters of the World.
Biogeography
Early studies of rotifer biogeography were dominated by
the certainty that all species could be dispersed worldwide,
an idea supported by the fact that the diapausing embryos
of monogononts and the xerosomes and xeroova of bdelloids may be transported by insects, birds, and mammals
(zoochory) and wind (anemochory). (See the later sections
on “Reproduction and Life History” and “Environmental Physiology: anhydrobiosis”) Such passive dispersal, it
was argued, is very effective at ensuring that most species
become globally distributed. However, data have accumulated suggesting that this conclusion is mistaken. In fact,
according to the last comprehensive survey of the literature
(Segers, 2007), few species have a cosmopolitan distribution (Figure 13.6). Therefore, rotifers may not be as easily
dispersed as previously thought. Moreover, once a dispersal stage arrives at a site, local abiotic conditions (e.g.,
pH, salinity, temperature) may be unsuitable for hatching.
Additionally, successful colonization requires hatchlings
compete for resources against competitors that have already
become adapted to the idiosyncrasies of the habitat. This
concept has been termed the Monopolization Hypothesis
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Phylum Rotifera
FIGURE 13.4 Variation in morphology of bdelloid rotifers. Scale
bars ca. 50 μm. Scanning electron photomicrographs courtesy of Diego
Fontaneto and Giulio Melone, University of Milan, Italy.
(a)
(d)
(b)
(e)
(De Meester et al., 2002). Thus, given the limits on dispersal and colonization, we should not be surprised that rotifers
show biogeographical patterns.
Recognizable trends in rotifer biogeography are evident. The fauna of Central America and the southern United
States have close affinities to tropical South American
fauna. Farther north, affinities with Europe are apparent,
and endemism is strong in Keratella, Notholca, and Synchaeta. Yet, no single hypothesis best explains what we currently know about the biogeography of rotifers. Depending
on the region, the rotifer fauna probably reflects the combined effects of recent glaciations and aridification, coupled
with subsequent zoochory and anemochory.
Nevertheless, there are some confounding issues to
studying patterns of rotifer distribution. Biogeographical
analyses are typically accomplished by collating information
collected from published works. Thus, one obvious problem
to achieving a comprehensive view of rotifer distribution is
that rotifer researchers have not explored everywhere. For
example, ∼52% of the species that have been recorded from
only one realm have been reported in the Palearctic, and
most of those from Europe. On the other hand, only ∼6% of
single-realm records have been reported in the Afrotropical
realm, which includes sub-Saharan Africa. These results are
not surprising, given that Europe is the birthplace of rotifer research. Thus, the pattern of distribution illustrated in
­Figure 13.6 probably reflects where the collections were
(c)
(f)
FIGURE 13.5 Variation in morphology of monogonont rotifers. (a) Asplanchna (foot absent), (b) Euchlanis (short foot with toes), (c) Epiphanes (prominent foot with toes), (d) Lecane (animal contracted into the lorica; short foot with prominent toes), (e) Testudinella (telescoping foot contracted into body),
(f) Cephalodella (animal somewhat flattened laterally by the preparation; toe prominent).
230
FIGURE 13.6 Occurrences of rotifers among the eight biogeographical realms. (Data compiled from Segers (2007).) Most species have been
found in a single biogeographic realm. Yet the question remains: Is this a
matter of uneven distribution of species (i.e., a biogeography) or vastly
uneven sampling or both?
made more than the actual distribution of species (Fontaneto et al., 2012). A second problem is that some groups,
such as bdelloids, are particularly u­nderstudied—some
rightfully say nearly ignored.
A separate issue is the phenomenon of seasonal polymorphism, which, in the past, confused workers who considered each morphotype to be a new species. A further
question that complicates these studies is the dilemma
of what actually constitutes a rotifer species; application
of the biological species concept is problematic because
males have never been described for many taxa. In practical terms, rotiferologists have been applying morphological criteria from the beginning of their discipline,
but the morphospecies concept has its own issues. Chief
among these is the fact that there are limited opportunities for workers to receive adequate training in rotifer
identification. Moreover, several workers have shown
that cryptic speciation is an important phenomenon in
rotifers, including species in the genera Brachionus, Epiphanes, Keratella, Notholca, Philodina, and Synchaeta.
Thus, minor variations in morphology, physiology, and/
or behavior that may be overlooked in identification may
actually reflect differences important enough to warrant
a new species designation. Efforts to clarify these matters will be greatly aided by modern genetic techniques,
which include microsatellite analysis and sequencing
molecular markers (e.g., CO1). Improved resolution of
species identity is essential to a complete understanding
of rotifer biogeography, a topic that Segers (2008) has
explored in detail.
Evolutionary Relationships
The phylogenetic position of rotifers is still an open question. The current view suggests that rotifers are somehow
related to the phylum Platyhelminthes, having been derived
either from an ancestral flatworm or as a sister group to
SECTION | III
Protozoa to Tardigrada
the flatworms. Another issue regarding the evolution of the
Rotifera is the question of its relationship to the parasitic
phylum Acanthocephala. The presence of the intracytoplasmic lamina (ICL) within a syncytial epidermis in both taxa,
along with similarities in the ultrastructure of their spermatozoa, suggests that acanthocephalans and rotifers are
closely related, even though acanthocephalans are 5–1500
times larger than the largest rotifer. Furthermore, detailed
molecular analysis aligns acanthocephalans firmly with the
rotifers, but exactly how they are related remains in doubt.
Thus, some researchers argue that similarities in structure
along with molecular evidence indicate that these two taxa
are sufficiently related to be recognized as forming their
own clade, the Syndermata (Sørensen and Giribet, 2006).
Within this clade, several possible permutations of rotifer
phylogeny have been postulated, including the following:
acanthocephalans are: (1) highly modified bdelloids; (2)
related to the group bdelloids + monogononts; (3) related to
the group seisonids + bdelloids; or (4) a sister group to all
rotifers.
Nonetheless, for the present, taxonomists in both
fields have retained the names Acanthocephala and
Rotifera, ignoring the question of their phylogeny.
Ultimately, integrative studies using total evidence
(morphological and molecular data) should help to
resolve issues of phylogenetic affinity. This goal is
made more feasible perhaps by studies of the transcriptome of Brachionus plicatilis Müller, 1786
and discovery that the mitochondrial genome of this
species is composed of two circular chromosomes of
unequal copy number. Indeed, much remains to be
accomplished, including a detailed analysis of a proposal
to group a suite of small, jawed taxa—Gnathostomulida,
Micrognathozoa, and Acanthocephala + Rotifera—into
a superphylum Gnathifera (G., gnath, jaw and L., fera,
bearing). Of course, to determine the phylogenetic relationships among these taxa, more molecular and morphological data, as well as a better sampling of taxa will be
necessary.
Evolution of bdelloids offers its own problems, but
research has supplied some exciting new insights into these
remarkable animals. Chief among the issues in bdelloid
evolution are the following: How did asexuality arise in
this group; how did their ability to become anhydrobiotic
appear; and how are deleterious mutations eliminated from
the population? Answers to these questions probably lie in
the fact that bdelloids have undergone genome duplication,
with the resulting gene copies evolving independent functions (i.e., they are degenerate tetraploids), as well as horizontal gene transfer, which has resulted in the acquisition
of a diverse array of genes from viruses, bacteria, and other
Metazoa (Ramulu et al., 2012). Current thinking on the evolution of the bdelloids and monogononts is that these two
groups separated at least 100 million years ago.
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Phylum Rotifera
GENERAL BIOLOGY
External Morphology
Rotifers exhibit a wide variety of morphologies; they are
saccate to cylindrical in shape, sometimes appearing wormlike. Typically, the body is composed of three or four
regions: head (corona), neck (in some forms), body (trunk),
and foot, although the foot often is absent in planktonic
forms. Although folds in the body may mark these regions,
they do not represent segments; thus rotifers are not metameric (segmented). In many species, especially bdelloids,
these creases function like joints or allow the body to collapse telescopically, thereby bending or shortening the animal, respectively (Figure 13.4). The generalizations noted
here do not describe all rotifers well. In some forms, the
neck and foot may be quite prominent, whereas in others
they are absent. When present, the foot often extends from
the body ventrally (Figure 13.1–13.3, 13.5). It usually possesses two toes, but this number may vary from 0 to 4. The
foot also may possess pedal glands the ducts of which exit
near the toes. These glands secrete an adhesive for temporary attachment to surfaces. On the other hand, juveniles of
sessile rotifers release cement that forms a bond with the
substrate that is not easily detached. If dislodged, the adults
of sessile species do not reattach.
In the subclass Monogononta, male rotifers (Figure
13.2) and juveniles of sessile rotifers (Figure 13.7) usually
are much smaller and have a morphology different from that
of adult females (Wallace, 1980; Wallace et al., 2006). In
addition, male rotifers are often structurally simpler (see
below: “Reproductive Systems”). Whenever males or juveniles are found in samples, their strikingly different morphologies may lead to improper identification.
Rotifers possess a body wall (integument) containing a
filament layer of varying thickness called the intracytoplasmic lamina (ICL) (Wallace et al., 2006; Wallace and Smith,
2009). This feature is shared only with the Acanthocephala,
and thus is thought to be synapomorphic. The ICL of B.
plicatilis, a halophilic rotifer, is composed of two filamentous, keratin-like proteins (Mr 39,000 and 47,000 Da) crosslinked by disulfide bonds (Wallace et al., 2006). Species in
which major portions of the integument are strengthened by
a thick ICL are termed loricate, and the thickened body wall
is simply referred to as the lorica (L., armor). The integument of species without an extensive ICL remains thin and
flexible. These rotifers are termed illoricate (L., il, without).
In general, thickness of the body wall is of little taxonomic
significance because extremes may be found within a single
family or genus (e.g., Cephalodella). Even in loricate species, regions of the integument possess fewer ICL filaments,
making the body wall flexible in that area. For example,
flexibility is found in the coronal field, in the foot or at its
junction with the body, and at articulations between movable spines and the trunk. In some rotifers, the surface of
(b)
(a)
(c)
(d)
FIGURE 13.7 Adults and juveniles (larvae) of some sessile species.
Family Flosculariidae: colony of adult Lacinularia flosculosa (a) and a juvenile somewhat compressed by the cover glass (b); Family Collothecidae:
solitary adult (c) and a newly hatched juvenile. Bars ∼100 μm. (a) and
(b), R.L. Wallace; (c) and (d), courtesy of Rick Hochberg, University of
Massachusetts at Lowell.
the body has various small projections (e.g., minute bumps
or ridges). Many rotifers possess assorted fixed or movable
spines, usually at the anterior and posterior ends. These serve
as protection from predators. The morphology of rotifers is
being better resolved by applying the techniques of electron microscopy and confocal laser scanning microscopy,
coupled with immunohistochemistry (Hochberg, 2009).
Some rotifers secrete materials from their body wall that
congeals into a gelatinous mass in which various items become
embedded, including inorganic silt, organic debris, bacteria,
algae, fecal pellets, and pseudofecal pellets. These materials
may form a tube that obstructs one’s view of the inside, but in
other species the jelly remains transparent. In a few species,
the secretions harden into a firm tube, as in the sessile monogonont genus Limnias, but in others the gelatinous material
hardens into a series of thin, loosely fitted tiles or plates, as in
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232
the bdelloid Mniobia incrassate (Murray, 1905). In some colonial species that produce tubes, the jelly mass coalesces into a
continuous matrix that surrounds all members of the colony.
On the other hand, the secretions of some bdelloids form small
amorphous tubes, termed nests (Habrotrocha and Rotaria) and
a few monogononts secrete only a thin jelly coat (Notommata
copeus Ehrenberg, 1834). Many workers suggest that tubedwelling rotifers do so to reduce predation, but the tube may
serve other functions. Ascomorpha eucaudis (Perty, 1850) and
Cephalodella forficula (Ehrenberg, 1832), for example, feed
on bacteria and/or algae that colonize the walls of their secretions, and the nests of bdelloids may serve as a way to retain
water during periods of dryness. A few species do not produce
tubes, but occupy cavities produced by other organisms: e.g.,
shells (tests) of ameba (Arcella and Difflugia), retort cells of
Sphagnum moss, and the lobules (cup-shaped, ventral leaf) of
the liverwort Frullania eboracensis.
Organ System Structure and Function
Internally, rotifers possess a perivisceral cavity commonly
termed the pseudocoelom, which bathes muscles and nerves,
and digestive, reproductive, and protonephridial organs
(Figures 13.1–13.3). (Note: We use the term pseudocoelom
for convenience, recognizing that some researchers recommend its elimination, as it is a polyphyletic condition.)
Respiratory and circulatory systems are absent in rotifers.
Although the characteristics noted above are unremarkable,
rotifers do possess two curious features. First, the cells of
all post-embryonic tissues are syncytial, i.e., multinucleate.
Second, all individuals of a species have a consistent number of nuclei in each organ throughout life. For example, citing an older work, Libbie Hyman noted that there are about
900 nuclei per female in Epiphanes senta (Müller, 1773)
and that the total number of cells in all species that had been
studied to date was between 900 and 1000 (Hyman, 1951).
Consistency in cell number, called eutely, is seen in a few
other invertebrates (e.g., nematodes, Chapter 14). Nuclei
may be seen using a standard light microscope, but special
optics such as differential interference contrast (Nomarski)
enhances their visibility.
Research on rotifer anatomy has proceeded rapidly during the past 30 years and promises to continue to increase
(e.g., Clément and Wurdak, 1991; Hochberg, 2006;
­Hochberg and Ablak Gurbuz, 2008; Hochberg, 2009; Smith
et al., 2010). However, no single work provides a comprehensive examination of rotifer structure; here we provide an
overview of the literature.
Corona
The classic description of the corona in rotifers is in the
form of two ciliated rings called the trochus and cingulum
(L., trocho, wheel and L., cingulum, belt). Nevertheless,
Protozoa to Tardigrada
there is considerable variation in the shape of the anterior
end, and a dozen or so different coronal forms have been
described based on overall shape, placement of the mouth,
and distribution of cilia. The cilia often are responsible
for production of water currents that are used in locomotion and feeding. In some forms, a prominent ciliated food
groove lies between the trochus and cingulum; this is best
seen in species of the order Flosculariacea. Yet, it is inadvisable to rely solely on coronal morphology in making identifications. For one thing, the head often retracts when the
animal is disturbed, and in fixed specimens the shape of
the corona can become distorted unless special methods of
preservation are used.
However, not all rotifers possess a ciliated corona.
Adults of the families Atrochidae and Collothecidae
exhibit the most extreme variation from the typical plan.
In collothecids, ciliation may be sparse, and long setae
surround the rim of a funnel-shaped structure known as
the infundibulum (L., a funnel) (Figure 13.8). These setae
prevent the escape of prey as the edges of the infundibulum fold over the victim, capturing it in a fashion similar to the Venus flytrap. Adult atrochids possess no setae
on their corona (Figure 13.9). In this small group, prey
are captured by a large infundibulum the edges of which
draw inward, capturing the prey like a purse net. Collothecids also possess a vestibulum, a small chamber located
beneath the infundibulum; here the prey is held before
being processed by the mastax (Figure 13.8). In other rotifers, ciliation may be limited to a ventral field or to a few
lobes, as is seen in some creeping monogononts and in
some bdelloids (Melone and Ricci, 1995). Other structures
that may be present on the corona include cirri, sensory
antennae, and palps.
Trophi and Gut
Food is processed by the mastax (a modified pharynx),
which is lined by a chitinous material developed as a set of
translucent jaws or trophi. Trophi work the food in various
ways (e.g., grasping, piercing, grinding, scraping) before
passing the food through the esophagus to the stomach. In
atrochid and collothecid rotifers, a portion of the mastax
is enlarged into a food-storage organ termed the proventriculus. The mastax leads into the esophagus and in turn
into the stomach. Most species possess an intestine and
anus, but the gut ends in a blind stomach in some genera
(e.g., Asplanchna, Asplanchnopus). The posterior portion
of the intestine receives eggs from the oviduct and fluid
from either a bladder or directly from paired protonephridia
(Figures 13.1–13.2). This part of the gut is termed a cloaca. Often the gut is pigmented, depending on the nature
of recently ingested material. Different species found in
the same sample may possess guts that vary in color due to
dietary preferences.
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Phylum Rotifera
Rotifer trophi are composed of several hard parts and
associated musculature that articulate in a specific spatial
arrangement. In their basic form, trophi consist of three
functional units: an incus (L., anvil) and paired mallei
(L., hammers) (Figure 13.10). The incus is composed of
three pieces: a fulcrum and a pair of rami (L., branches). The
latter move like forceps and articulate with the fulcrum at
their bases. Each malleus consists of two parts: manubrium
and uncus. The manubrium (L., handle) resembles a clubshaped structure extended at one end into a cauda and flared
at the other end (head). The manubrium articulates with a
toothed structure called the uncus (L., hook). The plane
of movement of the pieces comprising the malleus is at a
right angle to that of the rami. In some species, the trophi
may be modified by reduction of the basic parts, addition of
accessory structures, or asymmetrical development of one
or more of the pieces.
Trophi are important features in rotifer taxonomy,
as some classes, orders, families, and even species can
be determined based on details of the trophi alone. Nine
(a)
(c)
FIGURE 13.9 Cupelopagis vorax attached to the undersurface of an
aquatic plant. Members of Family Atrochidae possess no setae on their
corona.
(b)
(d)
FIGURE 13.8 Representatives of the order Collothecacea. (a) Collotheca trilobata, a sessile species; (b) Collotheca libera with one embryo, a planktonic species; (c) Collotheca ferox, a sessile species; (d) Collotheca sp., with one diapausing embryo.
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234
FIGURE 13.10 Malleate trophi of Cyrtonia tuba. In this scanning electron microscopy photomicrograph, all elements have been color-coded
to the labels. Photomicrographs courtesy of Diego Fontaneto and Giulio
Melone, University of Milan, Italy.
(a)
(b)
(d)
different types of trophi are recognized based on the size
and shape of the seven pieces and presence of any accessory
parts (Figure 13.11); transitional and aberrant types also are
known. The “Rotifer trophi Web page” (http://users.unimi.
it/melone/trophi/) provides an excellent overview of the different types of rotifer trophi.
In Malleate trophi (Figure 13.11(a)–(c),(e)–(f),(p)), all
parts of the incus and mallei are well developed and functional, but the rami are characteristically massive and may
possess teeth along the inner margin. Furthermore, the unci
have four to seven large teeth. This form works by grasping food and grinding it before pumping the crushed material into the esophagus. Malleate trophi are present in such
common rotifers as Brachionus, Keratella, and Lecane.
Malleoramate trophi (Figure 13.11(i)) are found only in
the order Flosculariacea and resemble the malleate form,
(k)
(c)
(m)
(l)
(f)
(e)
(o)
(g)
Protozoa to Tardigrada
(n)
(p)
(h)
(r)
(q)
(s)
(i)
(j)
(t)
(u)
FIGURE 13.11 Rotifer trophi types. (a) Malleate trophi of Epiphanes, ventral; (b) malleate trophi (Epiphanes), ventral elevated; (c) malleate trophi
(Epiphanes), lateral; (d) virgate trophi of Notommata, dorsal; (e) malleate trophi of Proales; (f) hypopharynx muscle in lateral aspect (Proales); (g)
incudate trophi of Asplanchna; (h) uncinate trophi of Collotheca; (i) malleoramate of Ptygura; (j) forcipate trophi of Dicranophorus; (k) asymmetrical
virgate trophi of Trichocera rattus; (l) virgate trophi of Eothinia; (m) virgate trophi of Itura, oral plates enlarged; (n) virgate trophi of Synchaeta with the
powerful hypopharynx muscle; (o) virgate trophi of Ascomorpha; (p) malleate trophi of Proales gigantea, (somewhat intermediate between malleate and
virgate types); (q) virgate trophi of Cephalodella, dorsal; (r) incudate trophi of Asplanchna; (s) virgate trophi of Cephalodella, lateral; (t) ramate trophi of
the bdelloid Pleureta lineata; (u) Cardate trophi of the family Lindiidae. Bars = 20 μm. (a)–(s) with permission from the authors Koste and Shiel (1987)
and CSIRO (http://www.publish.csiro.au/paper/IT9870949.htm); (u) with permission from the publisher Harring and Myers (1922).
Chapter | 13
Phylum Rotifera
except that, in the malleoramate form, the rami are strongly
toothed, and the unci possess many thin teeth. Similar to
the malleoramate form, ramate trophi (Figure 13.11(t)) have
large, semicircular shaped rami and unci with many teeth.
Ramate trophi lack a fulcrum and are limited to the subclass Bdelloidea. Only members of the order Collothecacea
possess uncinate trophi (Figure 13.11(h)). Although they
resemble the malleoramate, these trophi are characterized by unci possessing few teeth, usually with one large
one and a few small ones. Virgate trophi (Figure 13.11(d),
(k)–(o),(q),(s)) are modified for piercing and pumping, and
they generally can be recognized by their long fulcrum and
manubria and by the presence of powerful hypopharyngeal
muscles. Some trophi of this form are asymmetrical. Virgate
trophi are found in the common genera Notommata, Polyarthra, and Synchaeta. Forcipate trophi (Figure 13.11(j)),
as the name implies, have an action like forceps whereby
the trophi are projected from the mouth to grasp prey that
are then swallowed. Forcipate trophi are limited to the family Dicranophoridae. Incudate trophi (Figure 13.11(g),(r))
are restricted to the family Asplanchnidae. They also function by grasping prey with a forceps-like action, but this
form has a morphology different from that of the forcipate
type; the rami are quite large and the mallei very small. The
mastax actually initiates prey capture by creating suction,
drawing the prey into the mouth. Once captured, the prey
is stuffed into the stomach with the aid of the trophi. Lacking an intestine and anus, species of the genera Asplanchna
and Asplanchnopus also use their trophi to extract indigestible materials from the stomach. Cardate trophi ­(Figure
13.11(u)) are found only in the family Lindiidae and function by producing a pumping action, without the hypopharyngeal muscle. The fulcrate type of trophi has been
described as an aberrant form and is incompletely understood. This form is found only in the class Pararotatoria (see
above), a very small group of marine rotifers comprising
only four species within two genera: Paraseison (1 species)
and Seison (3 species).
A relatively unexplored facet of rotifer nutritional physiology is digestive enzyme function. Some work has been
done with homogenates of whole animals (Kleinow and
Röhrig, 1995), but histochemical and enzyme-labeled fluorescence techniques have permitted researchers to locate specific enzymes within organs (e.g., Strojsová and Vrba, 2007).
235
known as body-wall outgrowths (Asplanchna, Figure 13.12).
In others, such as Brachionus calyciflorus Pallas, 1766
(Figure 13.13), increasing the pressure results in a flexing of
the articulation between spines and the body in the posteriolateral region of the animal. This causes the spines to swing
outward and stiffen. In this extended position, the spines
interfere with predators (e.g., Asplanchna) that would otherwise consume undefended prey. Striated, longitudinal muscles are responsible for retracting the corona and foot and for
moving certain articulating projections that are not moved by
changes in the hydrostatic pressure of the body cavity. Hexarthra, for example ­(Figure 13.14), possesses powerful muscles
that control movement of arm-like locomotory appendages.
Contractions of these muscles cause a swift sweep of the
arms that results in a rapid displacement or jump of the rotifer. Jumps in members of the genus Polyarthra are achieved
by the sweeping movement of paddle-shaped appendages.
However, in the genus Filinia, long setae (spines or bristles)
are moved by the action of an increase in the hydrostatic
pressure.
FIGURE 13.12 Body form variability in the genus Asplanchna sieboldii.
BWO = body wall outgrowths. Drawn from original photomicrographs
courtesy of John J. Gilbert, Dartmouth College.
Muscular System
The muscular system consists of small groups of longitudinal
and circular muscles inserted at various points on the integument or between the integument and viscera (Figure 13.2).
In loricate species, the integument provides a firm structure
against which the muscles work. In some species, muscles
retract the corona, which increases pressure within the body
cavity, thus expanding flaccid portions of the integument,
FIGURE 13.13 Induction of spines in Brachionus calyciflorus. Left
to right: spineless individual, three individuals with increasing spine
development, individual with fully developed spines. Figure modified from Gilbert (1967), with the kind permission of the author and
www.schweizerbart.de.
236
SECTION | III
Protozoa to Tardigrada
FIGURE 13.14 Hexarthra showing positioning of muscles that initiate
jumps. Bar = 100 μm.
Muscles also are present in the viscera, particularly
in the mastax and stomach. Staining rotifer musculature
with a fluorescent dye linked to phalloidan and examining
specimens with a confocal laser-scanning microscope has
revealed much finer details of these tissues (e.g., Hochberg
and Ablak Gurbuz, 2008).
FIGURE 13.15 Schematic representation of the nervous system of
Plationus patulus revealed with immunohistochemistry (anti-serotonin,
anti-FMRFamide, anti-neurofilament) and confocal microscopy. Note the
presence of individual neurons in the coronal ciliated field. Abbreviations:
br = brain; cnr = coronal nerve ring; ma = mastax; mg = mastax ganglion;
pg = pedal ganglion; pn = pedal neuron; st = stomach and associated gastric
neurons; tc = transverse commissure; vlc = ventrolateral nerve cord. Figure
courtesy of Rick Hochberg, University of Massachusetts at Lowell.
Neural System
The nervous system is simple, comprising only a cerebral
ganglion or brain (Figure 13.1) located dorsally on the
mastax, a few other ganglia present in the mastax and foot,
and three types of sensory organs: mechano-, chemo-, and
photoreceptors (Clément and Wurdak, 1991). Mechanoreceptive bristles are situated on the corona, whereas several
antennae are located elsewhere on the body, usually laterally and caudally (Figures 13.1–13.2). Chemoreceptive
pores also are present on the corona. Many species possess
one or more photoreceptive eyespots, sometimes containing
a red pigment. When present, eyespots are located at the
anterior end, usually near the brain. Although some rotifers
retain their eyespots throughout life, sessile rotifers often
lose them after the juveniles attach to a substrate. Paired,
ventral nerve cords proceed from the brain along the length
of the body into the foot. Several other ganglia are usually
found in the nerve cords at the exit points for lateral nerves
(Figure 13.15).
One interesting structure found in the apical region of
many bdelloid and monogonont rotifers is the retrocerebral
organ (RCO). This structure consists of paired subcerebral glands and an unpaired retrocerebral sac, both with
ducts that lead to the surface of the corona (Figure 13.1).
Although the RCO function is not completely understood,
fewer protonephridia occur in rotifers with well-developed
RCOs (Edmondson, 1959). Some researchers have speculated that the RCO functions as an exocrine gland perhaps
lubricating the rotifer’s apical end (Clément and Wurdak,
1991).
Information on neurobiochemistry remains limited, but
immunocytochemical studies have shown that acetylcholine functions as a neurotransmitter (a cholinergic system)
in species from at least six families. In addition, norepinephrine neuroreceptor sites (an adrenergic system) have
been reported in B. plicatilis, and widespread catecholaminergic neuronal systems have been observed in species
of Asplanchna and Brachionus. Research also has identified serotonin and FMRFamide immunoreactive neurons in
monogononts. Nicotinic receptors also have been identified
in monogononts and at least one bdelloid. Neuron size and
innervation varied among species, but neuron number was
constant and species-specific.
Excretory System: Protonephridium
A paired protonephridial system composed of tubules and
flame cells functions in excretion of nitrogenous wastes and
osmoregulation in all rotifers (Figures 13.1–13.2). Usually
there are only a small number of flame cells (fewer than
Chapter | 13
237
Phylum Rotifera
six), but large rotifers may possess many more. For example, Asplanchna sieboldii (Leydig, 1854) may have up to
100 flame cells (Ruttner-Kolisko, 1974). Normally, these
tubules drain into a urinary bladder (Figure 13.2) that leads
to a cloaca, but the bladder is absent in some species and a
contractile cloaca assumes its function.
Reproductive System
Gonads are paired in both marine seisonids (class Pararotatoria) and the bdelloids (class Eurotatoria, subclass Bdelloidea); but in the latter, males are completely unknown and
reproduction is by parthenogenesis. Monogonont rotifers
(class Eurotatoria, subclass Monogononta) have only one
gonad. Although males are present in this group, they have
not been described for a large number of species. Even so,
it is generally assumed that all monogononts can produce
males given the proper conditions, or at least that the ancestral forms were capable of male production. However, when
males are made the primary episode of sexual reproduction
typically lasts for a few days to a week. Thus, unless collections are made frequently, male rotifers may never be
observed. Nevertheless, even when a portion of a population is producing males, parthenogenetic reproduction usually continues in the remainder.
Sexual reproduction (mixis) is initiated in many monogononts when females respond to a crowding signal via a
labile, quorum-sensing molecule (mixis signal) released
into the environment by the females themselves; this results
in the production of males. Males may fertilize females,
which in turn produce embryos that do not immediately
hatch from their egg case; these are called diapausing
embryos (see the later section on “Reproduction and Life
History”). A few strains of the genus Brachionus have been
reported to be obligate parthenogens, apparently having lost
the capacity for sexual reproduction after long-term culture
in a chemostat.
The reproductive organs of females are composed of
three units: ovary, vitellarium, and follicular layer. The
ovary of a rotifer is a small, syncytial mass that is closely
associated with the yolk-producing vitellarium. At birth, the
adult complement of oocytes already has been formed in the
ovary. The vitellarium also is syncytial with a specific number of nuclei—a characteristic useful in the taxonomy of
some species. The follicular layer surrounds both the ovary
and vitellarium. In some species, this layer forms the oviduct, which connects to the posterior portion of the gut at
the cloaca (Figure 13.1).
In general, monogonont males are smaller, swim faster,
and have shorter life spans than females of the species, but
this generalization does not hold for all species (e.g., Rhinoglena; Melone, 2001). In those species with small males
(e.g., Asplanchna, Brachionus, Conochilus), the gut of the
male is reduced or is absent entirely (Figure 13.16). In these
(a)
(b)
(c)
(d)
(e)
(f)
FIGURE 13.16 Examples of male rotifers. (a) Anuraeopsis fissa
(bar = 10 μm); (b) Asplanchna girodi (bar = 100 μm); (c) Brachionus calyciflorus (bar = 20 μm); (d) Brachious plicatilis (bar = 50 μm); (e) Lecane
quadridentata (bar = 15 μm); (f) Platyias quadricornis (bar = 10 μm).
(Photomicrographs (a)–(c) & (e), (f) courtesy of Roberto Rico-Martínez,
Universidad Autónoma de Aguascalientes, Aguascalientes, México; d,
Modified, with permission, from Pourriot and Francez (1986).)
so-called dwarf males, the rudimentary gut serves as an
energy source (Ricci and Melone, 1998), but there are other
structural reductions as well (e.g., corona) (­Gilbert and
­Williamson, 1983). The single testis is large and saccate,
usually containing fewer than 50 freely floating mature
sperm. A ciliated vas deferens leads from the testis to the
penis, often with one or rarely two pairs of accessory (prostate) glands that discharge into it. In sessile species, the
male is free swimming, whereas the adult female remains
affixed to the substrate.
Most rotifers are oviparous, releasing their eggs outside
the body where the embryos develop. Many planktonic species carry their eggs attached to the female by a thin thread
(Brachionus), fix them to a substrate (Euchlanis), or release
them into the plankton (Notholca). The sessile species
­Sinantherina socialis (Linnaeus, 1758) carries its eggs on a
specialized structure called the oviferon, which is located on
238
its elongated foot below the anus. However, a few species are
ovoviviparous (e.g., Asplanchna and Cupelopagis), and thus
retain their embryos in the body until the offspring hatches.
Environmental Physiology
Locomotion
Most rotifers swim during at least a portion of their life
cycle, but those lacking a foot (e.g., Asplanchna, Keratella,
and Notholca) never attach to surfaces, even temporarily.
Swimming can influence acquisition of food and mates,
increase number of encounters with predators, and promote
dispersal of the juveniles of sessile forms. Some rotifers
are unusual in that they can swim, but routinely remain in
mucus tubes attached to a substrate (e.g., certain Cephalodella species), while others are free-floating in mucus
sheaths (Ascomorpha). In addition, some sessile species
have relatives that have evolutionarily abandoned the sessile existence becoming mobile; these include Collotheca
libera (Zacharis, 1894) (Figure 13.8(b)), Ptygura libera
Myers, 1934, and Sinantherina semibullata (Thorpe, 1893).
Rotifers often swim in a helical pattern, so that the actual
distance traveled is greater than the linear displacement
(Starkweather, 1987). However, for practical reasons, most
researchers do not attempt to calculate absolute distance
when considering the distance traveled, although automatic
tracking systems have made this tedious work easier. While
the theoretical power requirements of swimming (i.e., the
energy required to overcome water resistance) is <1% of
total metabolism, actual energetic cost appears to be much
greater. In B. plicatilis, this cost has been calculated to be
∼38% of total metabolism (Epp and Lewis, 1984).
Several researchers have measured the swimming
speeds of male and female B. plicatilis. Swimming speeds
also have been measured for other species such as Anuraeopsis fissa Gosse, 1851, Asplanchna brightwellii Gosse,
1850, B. calyciflorus, Euchlanis dilatata Ehrenberg, 1832,
and Keratella americana Carlin, 1943. Collectively, these
studies show that swimming speed is temperature dependent
and varies with species and strain, age, number of eggs carried, food levels, and body mass. Values recorded for B. plicatilis at 25 °C range from 0.5 to 1.5 mm/s for females and
1.3–1.5 mm/s for males, with young and old females swimming about 30% slower than mature females. Asplanchna
brightwellii and Keratella spp. females normally swim at
0.9 and 0.5 mm/s, respectively. Swimming speeds ranging from 0.17 to 0.54 mm/s were reported for 11 species
of freshwater rotifers by Santos-Medrano et al. (2001). In
this study Reynolds numbers varied between 0.023 and
0.301, with drag coefficients ranging from 9.7 × 10−7 to
1.6 × 10−5 N. We can infer that at these low Reynolds numbers the watery world is governed by laminar flow: i.e., at
their size and swimming speed there is no turbulence.
SECTION | III
Protozoa to Tardigrada
Little is known about the swimming speeds of the juveniles
of most sessile rotifers, but swimming speeds for juveniles of
Ptygura beauchampi Edmondson, 1940 are known to vary with
age. For example, newborns up to mid-aged juveniles (∼3 h
old) swam at about 2–2.5 mm/s. This speed is faster than the
rates reported for B. plicatilis males. However, older juveniles
(>4.5 h) swim at ∼1.0 mm/s. Concomitant with these changes
in swimming speed is an increase in turning frequency.
Correlations of rotifer ultrastructure with swimming
speed, turning frequency, and other behaviors have been
well studied and have demonstrated that rotifers are excellent models for comparative neurobehavioral studies. For
example, swimming speed, mean turning angle, and angular
speed varies in B. calyciflorus as a function of the condition of the medium and animal nutritive state. Nevertheless,
much more work remains to be done before a complete synthesis of structure, function, and behavior will be possible.
Spines and other appendages influence both swimming
speed and sinking rates in rotifers. For example, unspined
Keratella testudo (Ehrenberg, 1832) generally swim faster
and sink more slowly than spined forms. On the other
hand, Polyarthra normally swims at a much slower velocity (0.24 mm/s) than any species previously discussed, but it
is capable of very short bursts (about 0.065 s long) of rapid
movements called jumps (see above). During a jump, Polyarthra may attain velocities greater than 50 mm/s, with a
mean velocity of 35 mm/s, or ∼100 times its normal swimming speed (Starkweather, 1987). Jumps are produced by
movements of 12 appendages, called paddles, that articulate
with the body near the head. When Polyarthra detects a disturbance in the water up to a few body lengths away, powerful striated muscles rapidly flex some of the paddles upward,
initiating the jump. Then, during the jump, the paddles
return to their original positions, and the rotifer is displaced
an average of 1.25 mm (ca. 12 body lengths). Jumps help
this rotifer escape invertebrate predators, such as the rotifer
Asplanchna and first instars of the phantom midge Chaoborus, as well as the filtering currents of microcrustaceans
such as Daphnia. They likewise are effective against naive
workers attempting to remove Polyarthra from plankton
samples using pipettes! Hexarthra also can jump rapidly,
but movement in these forms has not been as well studied.
In contrast, Keratella cannot jump to avoid the filtering currents of large daphnids; thus, the animals are severely damaged or killed when swept into the branchial chambers of
cladocerans. However, Keratella is not without some escape
abilities; it can increase its swimming velocity by a factor of
about 3.5 when it encounters inhalant currents of Daphnia
or comes into contact with the predatory rotifer Asplanchna.
Physiological Ecology
Physiological tolerances of organisms prescribe environments where survival and reproduction are possible. Thus,
Chapter | 13
Phylum Rotifera
an environmental tolerance curve for a species summarizes
the range of environments where reproduction occurs and
is bounded by the upper and lower lethal limits for the species. Within a tolerance curve the environmental optimum
for a species is that environment where survival and reproduction are maximal. Therefore, a set of tolerance curves
(including temperature, pH, salinity, oxygen concentration,
etc.) indicates the breadth of adaptation of a species and its
niche dimensions.
Environmental tolerances and niche widths have been
characterized for few rotifer species. For example, Epp and
Lewis (1980) described the response of B. plicatilis to temperature. Their study determined respiration rate over temperatures ranging from 15 °C to 32 °C, and recorded Q10s of
1.9–2.4 for a broad temperature interval. Respiration levels
off in the range of 20–28 °C, indicating that B. plicatilis can
maintain a constant metabolic rate over this temperature
range. At higher and lower temperatures, respiration rate
increased, presumably because of thermal stress beyond the
homeostatic capability of this species. Snell (1986) showed
that amictic and mictic females have different temperature
tolerance curves. Amictic B. plicatilis females reproduced
at 20 °C and 40 °C, whereas mictic females did not. In general, amictic females reproduced over a broader environmental range of temperature, salinity, and food levels than
did mictic females.
Effects of pH on distribution and abundance of rotifers is a topic that has received a good deal of attention.
However, because hydrogen ion concentration is related
to other important chemical parameters in freshwaters,
the effect of pH on rotifer occurrence may be indirect.
Nevertheless, some extensive early work by F. Myers in
the 1930s demonstrated that rotifers could be classified
into a few broad groups based on pH alone: alkaline species, acid species, and those with a broad range. In the
late 1980s, B. Bērziņš and B. Pejler, working from a large
number of habitats in Sweden, concluded that species
found in oligotrophic waters had pH optima at or below
neutrality, whereas those species common to eutrophic
waters had optima at or above neutrality. Moreover, they
noted that acid-water species were often nonplanktonic
or semiplanktonic. Such generalizations are, of course,
never exact, and other factors are important. For example,
Weithoff (2004) documented occurrence of two planktonic
species—Cephalodella hoodii (Gosse, 1886) and Elosa
worallii Lord, 1891—in an acid lake (pH 2.7) in Germany,
where temperature and resource availability contributed to
niche separation. Much less is known of the specific metabolic responses of rotifers to pH, but in B. plicatilis swimming activity and respiration rate does not significantly
vary at pH values of 6.5–8.5 (Epp and ­Winston, 1978).
However, swimming was reduced below pH 5.6 and above
pH 9, with alkaline waters depressing swimming activity
more than acidic conditions.
239
Regardless of the difficulties of interpreting the effects
of pH on rotifers, we know from the work of Frost et al.
(2006) that a shift in pH can radically alter zooplankton
community structure. For example, increasing acidification of lakes as a result of acid precipitation has been widespread in North America, and this has led to changes in the
composition of the zooplankton, e.g., increasing dominance
of Keratella taurocephala Myers, 1938. This observation
has been repeated in lakes that have been experimentally
acidified. In general, it appears that rotifers become more
important as members of the zooplankton with increasing
acidity.
Individual rotifers rarely experience large fluctuations
in osmotic conditions over their relatively short lifetime.
However, a population living in a small water body subject
to periodic inundation followed by evaporation may need
to cope with increasingly greater solute concentrations.
These habitats include tide pools, marine coastal ponds, and
desert rock pools. Brachionus plicatilis, a common euryhaline species, survives osmolarities over the range of two
to >95 psu (i.e., ∼50 to >2750 mOsmol/l). Transferring of
B. plicatilis directly from ∼1.5 to ∼33 psu causes considerable mortality, but acclimation to elevated osmolarities
can be achieved by gradually increasing ionic concentrations. On the other hand, B. plicatilis does not tolerate low
osmolarities well; this probably accounts for its restriction
to alkaline and brackish waters. This well-studied species
can probably tolerate these extreme salinities because it can
regulate its internal bodily fluid via a membrane-bound,
Na+/K+ ATPase pump (Lowe et al., 2005).
Although most rotifers require oxygen concentrations
significantly above 1.0 mg/l, some tolerate anaerobic or
near anaerobic conditions for short periods. Other species
routinely live in oxygen-poor regions, such as the hypolimnion of eutrophic lakes or in sewage ponds.
Rotifer resistance to starvation has been investigated in
a number of species. For example, in the bdelloid Macrotrachela quadricornifera Milne, 1886 survivors of a long
starvation period (20–60 days) resumed reproduction after
being fed (Ricci and Perletti, 2006). Life history parameters
like mean lifespan and lifetime fecundity were not different
in starved bdelloids after the resumption of feeding, suggesting that starved rotifers simply shift their schedule of survival
and fecundity. Thus, in bdelloids, metabolic reduction during
starvation is similar to their physiological response to drought.
Using life table experiments, Weithoff (2007) investigated
the response of two monogononts, Cephalodella sp. and
E. worallii, to starvation. Using food concentrations below the
threshold for population growth, he found a tradeoff only in
E. worallii, which increased lifespan at the expense of reproduction. Differential resistance to starvation is important, as
it can determine the outcome of competition and thus influence community structure. For example, research has shown
that starvation resistance was higher for Keratella cochlearis
SECTION | III
240
(Gosse, 1851) grown on N-limited Cryptomonas rather than
nutrient-sufficient algae.
Food quality also has been shown to explain the vertical
distribution of Cephalodella sp. and E. worallii in a meromictic lake in Germany (Weithoff and Wacker, 2007). Laboratory
research confirmed that fecundity, growth, and resistance to
starvation by E. worallii were negatively affected when the
rotifer was fed the alga Chlamydomonas acidophilia that
had been grown under heterotrophic vs. autotrophic culture
conditions. However, Cephalodella sp. grew on a diet of
C. acidophilia regardless of conditions under which the algae
had been grown. In the field, a population of E. worallii was
abundant in the epilimnion (above 6 m), where individuals of C. acidophilia would have been growing autotrophically. On the other hand, individuals of Cephalodella sp.
were more abundant in the hypolimnion (below 6 m) where
C. acidophilia would have been growing heterotrophically.
Environmental Toxicology
Because rotifers fill an important ecological role and are
relatively easy to culture, they have been important in
assessing the toxicity of chemicals in the laboratory, as well
as examining the effects of pollutants in natural habitats.
The response of rotifers to a variety of toxicants has
been characterized in both natural and laboratory populations. Most methods have used brachionid rotifers, but other
species have also been used. These involve both short-term
(acute) and long-term (chronic) toxicity tests. The test
materials include crude oil and other petrochemicals, oil
dispersants, detergents, free ammonia, wastewater, endocrine disruptors, anti-infective agents, cyanotoxins, insecticides, herbicides, and heavy metals. Research has shown
how exposure to these factors modifies rotifer life table
parameters such as age-specific survivorship and fecundity,
and the production of diapausing embryos.
Standardization of toxicity tests provides a way to compare the effects of toxicants, which may otherwise differ
depending on the protocol. A standardized acute toxicity test
has been described that employs freshwater (B. calyciflorus)
and marine (B. plicatilis) test animals (ASTM, 2012). This
procedure is less expensive than many others because it
uses test animals hatched from diapausing embryos, so it
is not necessary to maintain stock cultures. Moreover, the
test is simple, rapid, and sensitive. In addition, a standardized chronic toxicity test based on asexual reproduction has
been published in Standard Methods for the Examination of
Water and Wastewater (Snell, 1998). Because toxic agents
affect swimming and feeding behaviors, these phenomena
also have been used to assess toxicity. Besides such direct
effects, toxicants can modify rotifer community structure
through indirect effects, for example by altering food composition and thereby changing the outcome of resource
competition.
Protozoa to Tardigrada
In general, rotifers seem to serve as good indicators of
water quality in natural environments. In Europe, for example, composition of the rotifer community has been advocated as a component in the Saprobic Index. (The Saprobic
Index is a measure of saprobity, an estimate of the level of
organic pollution as measured by a combination of the biological oxidation demand of a water sample and the presence of specific indicator organisms in the habitat.)
Studies of natural and experimental contamination of
water bodies have shown that rotifer species composition
and abundance change depending on the toxic agent and
its concentration. In a study of the effects of artificial acidification on a lake in north central Wisconsin (USA), Frost
et al. (2006) showed that whereas the zooplankton community (including rotifers) improved after release from the
acid stress, the trajectory of the recovery followed a different path from the decline, thus indicating a substantial
hysteresis. Even a temporary contaminant can have a strong
effect on rotifers. After a spill of the so-called Red Sludge in
the Danube, the rotifer community disappeared. Although
it recovered a few weeks later, the levels were lower than
before the spill (Schöll and Szövényi, 2011). Sustained,
heavy contamination can have a profound effect on zooplankton composition. In a reservoir in the Chelyabinsk
region of Russia, which appears to be the most radioactively
contaminated inland waters in the world, the plankton community is composed of a near monoculture of cyanobacteria
and rotifers (Pryakhin et al., 2012).
Anhydrobiosis
The ability of rotifers to tolerate desiccation and then be
revived later has been known since the early 1700s, when
Leeuwenhoek described rehydration of rotifers present in
sediments collected from dry rain gutters. This phenomenon, called anhydrobiosis, is known to occur in both adults
and embryos of many bdelloids. However, in monogononts,
dormancy occurs only in the diapausing embryos, which
also are capable of anhydrobiosis. These embryos are the
product of sexual reproduction (see the section “Reproduction and Life History: Diapausing Embryos in Sediments”).
While in the desiccated state, bdelloids and diapausing
embryos are capable of passive dispersal by air currents
(anemochory) or by mobile animals (zoochory), either
attached on the outside (ectozoochory) or in their digestive
tracks (endozoochory).
An anhydrobiotic adult bdelloid rotifer resembles a
wrinkled barrel and has been called a tun (Middle English,
meaning barrel); unfortunately, this descriptive term is also
routinely applied to anhydrobiotic tardigrades (Chapter 17).
To avoid confusion, the term xerosome (G., xero, dry and
G., soma, body) has been proposed (Wallace and Smith,
2009) for anhydrobiotic adult bdelloids. In xerosomes,
the head and foot retracts into the animal’s trunk; at this
Chapter | 13
Phylum Rotifera
time, the female may deposit a mature egg (Figure 13.17).
An egg, really an embryo, deposited during the desiccation process also can withstand desiccation and is termed
a xerooum (G., dry egg). While the desiccation process is
complex, the adaptive potential is substantial, as many bdelloids inhabit environments that dry completely at irregular
intervals (Ricci et al., 2007). A few strictly aquatic bdelloids
cannot withstand desiccation.
Bdelloid anhydrobiosis involves more than simple drying; unless loss of metabolic water proceeds slowly, the
rotifer usually dies. During anhydrobiosis, the fine structure of cells is retained, but in a greatly modified state.
Changes that occur internally include at least a 50% reduction in the volume of the body cavity, a condensation of
cells and organs, and a decrease in cytoplasmic volume, so
that the entire animal is only about 25–30% of its original
size. Nuclei, mitochondria, endoplasmic reticula, and other
organelles form a compact mass within cells of the xerosomes (Ricci et al., 2007).
Xerosomes have been reported to be viable even after
more than two decades in the anhydrobiotic state. Recovery
from anhydrobiosis may require as little as 10 minutes, or
it make take several hours, according to prevailing environmental conditions. Survival is influenced by both biotic and
abiotic factors. For example, young and starved animals are
less likely to survive anhydrobiosis. Moist environments
and high temperatures during desiccation also reduce recovery. A noteworthy feature of anhydrobiosis in bdelloids is
that the time spent as a xerosome does not affect the total
life span; this phenomenon has been termed the Sleeping
Beauty Model (Ricci and Caprioli, 2005). For example,
Ricci and colleagues have shown that 8-day-old Adineta
ricciae Segers and Shiel, 2005 desiccated for 7 days lived as
long as a group from the same cohort of animals that was not
FIGURE 13.17 Adult bdelloid xerosome (left) and its desiccated embryo
(xerooum) (right). Scanning electron photomicrograph courtesy of Giulio
Melone, University of Milan, Italy.
241
desiccated (mean (±SD) = 41.2 (1.2) and 37.7 (1.2), respectively). However, the desiccated group did produce significantly more offspring per animal over their lifetime (mean
(±SD) = 37.6 (0.4) and 31.9 (0.4)), respectively. Given that
rotifers are eutelic, this increase in fecundity offers a mystery, especially as Ricci and colleagues noted that this species possesses only about 30 nuclei in its germarium. Ricci
and coworkers suggested that anhydrobiosis may stimulate
oocyte production by the germarium but then remind us that
this idea runs counter to the current understanding of eutely.
Further complicating our understanding of anhydrobiosis in
bdelloids is the fact that, in at least two bdelloid species
(A. ricciae and M. quadricornifera), repeated desiccations
and rehydrations appear to enhance fitness over those in
continuous culture.
Anhydrobiosis in bdelloids occurs without production
of the nonreducing disaccharide trehalose or an analogous
molecule, and no trehalose synthetase genes have been
identified (Mark Welch et al., 2009). Nevertheless, hydrophilic proteins found in desiccation-tolerant plants become
abundant late in the embryo of the common bdelloid
Philodina roseola Ehrenberg, 1832. Although trehalose is
absent from bdelloids, it is present in desiccated diapausing embryos of the monogonont B. plicatilis. The molecular
mechanisms of anhydrobiosis need further investigation,
and bdelloids seem to be an excellent research model for
dormancy studies. One intriguing proposition is that extra
copies of the gene for alpha tubulin may be related to their
ability to withstand desiccation. This is suggested because
11 to 13 copies have been found in bdelloids (Eyres et al.,
2012) when only four to eight would be expected in these
degenerate tetraploids.
Bdelloids also possess extreme resistance to 137Cs
gamma ionization radiation (IR) (Mark Welch et al.,
2009). For example the bdelloids Adineta vaga (Davis,
1873) and P. roseola reproduce (albeit at a reduced rate
over un-irradiated controls) even after exposure to doses
of more than 800 Gy (by comparison, a chest X-ray will
deliver ∼10−4 Gy). In contrast, the monogonont E. dilatata
experienced greater reduction of fecundity, but at a radiation dose of only 20% that of the bdelloids. Moreover,
although the bdelloids experienced hundreds of doublestranded breaks (DSB) to their DNA during exposure to
the radiation, the damaged was repaired. Thus, bdelloids
are not resistant to IR but possess an extraordinary ability to repair DSB. Their ability to tolerate desiccation and
ionization radiation may stem from the fact that bdelloids
are degenerate tetraploids (see above), with collinear pairs
of chromosomes that may serve as templates for DSB
repair via a form of homologous recombination (Mark
Welch et al., 2009). Moreover, processes associated with
DSB repair may foster removal of deleterious elements
and have contributed to the persistence of these ancient
asexual animals.
242
Generalized Stress Responses
Studies have shown that rotifers respond to stresses such
as heat, UV radiation, and low pH by producing an array
of heat shock proteins (HSP) (e.g., Smith et al., 2012).
In addition, Late Embryo Abundant genes (LEA) help in
maintaining homeo-osmotic conditions (Denekamp et al.,
2010). However, recent work indicates that rotifers also produce Transient Stress Granules (SG) in response to stress,
but the extent to which they are produced varies by the
nature and duration of the stress (Jones et al., 2013). Additional research will, no doubt, elucidate the relationship
among environmental stressors and these stress response
mechanisms.
ECOLOGY AND EVOLUTION
Diversity and Distribution
Phenotypic Variation
Phenotypic variation is an important adaptive mechanism
in rotifers, but has posed difficult problems for systematists. Intraspecific variation arises by several mechanisms
including cyclomorphosis, dietary- and predator-induced
polymorphisms, dwarfisms, and polymorphisms in hatchlings from diapausing embryos. Cyclomorphosis is the
seasonal phenotypic change in body size, spine length, pigmentation, and/or ornamentation found in successive generations of zooplankton. These changes are morphological
alterations in the individuals of a single population that are
related to physical, chemical, or biological features of the
environment. Each different morphological form is called a
morphotype (or morph). Specifically excluded from cyclomorphic changes are seasonal succession of sibling species
and clonal replacements of genotypes (both of which are
genetic changes in populations or communities), as well as
induction of spines or changes in shape due to the presence
of predators or specific biochemicals in the diet.
Gilbert (1980) described a striking phenotypic change
in morphology associated with a dietary polymorphism for
three Asplanchna species (A. brightwellii, A. intermedia
Hudson, 1886, A. sieboldii). Diets that include the plant
product α-tocopherol (vitamin E) induce saccate females, the
smallest morphotype, to produce cruciform daughters. Cruciforms have lateral outgrowths of the body wall ­(Figure 13.12)
that protect them from cannibalism by conspecifics by making them larger and thus more difficult to ingest if captured.
In the presence of vitamin E and certain prey types, cruciforms can produce a third morphotype called campanulates (more prevalent in A. sieboldii and A. intermedia).
Campanulates are very large females (>2000 μm) that heavily cannibalize saccate females. Female polymorphism is
much less pronounced in A. brightwellii; their increase in
body size is only 50–60% larger than the norm. Dietary
SECTION | III
Protozoa to Tardigrada
polymorphism in Asplanchna (gigantism) may have evolved
originally as a generalized growth response to larger prey
typical of eutrophic waters. The tocopherol response probably is adaptive, because it may signal availability of nutritious rotifer and microcrustacean prey.
Another important source of phenotypic variation is
predator-induced polymorphism. This topic was briefly
considered above (“Environmental Physiology: Locomotion”). Spined and unspined forms had been recognized
in several rotifer species for many years, but the causes
and consequences of these variations remained an enigma
­(Figure 13.13). Induction of spines was first demonstrated
in B. calyciflorus: the offspring of adults exposed to culture medium that had previously held predatory Asplanchna
developed spines that dramatically reduced their vulnerability to predation by Asplanchna. Additional research has
shown that several other prey species are capable of spine
induction by Asplanchna, as well as by other predators.
Two additional kinds of variation are aptera generations in Polyarthra (i.e., polymorphisms in the hatchlings
of diapausing embryos) and a body size that is smaller than
is commonly seem. The latter has been called dwarfism,
but this term should not be confused with the smaller size
often seen in monogonont males. Aptera morphotypes were
initially thought to be different species of Polyarthra, but
later were shown to be forms lacking the paddles that are
characteristic of this genus. Only the generation hatching
from the egg bank of diapausing embryos lacks paddles;
their parthenogenetic offspring develop into typical morphotypes with these appendages. Similar polymorphisms
between diapausing embryos hatchlings and parthenogenetic generations were described for Keratella quadrata
(Müller, 1786) and are suspected for Notholca acuminata
(Ehrenberg, 1832). Dwarfism in Brachionus caudatus
Barrois and Daday, 1894, as reported for crater lakes in
the Cameroon, is characterized by reduced body size and
spination compared to normal morphotypes; high temperature, combined with reduced food supply and a different
suite of predators, may account for this condition. However,
we cannot discount the possibility that these populations
actually represent cryptic species.
Another source of phenotypic variation in body size is
polyploidy. For example, Walsh and Zhang (1992) reported
substantial variation in body size for different populations
of E. dilatata. During a 12-month period of intensive sampling in Devils Lake (Oregon), these workers observed
two distinct body sizes: a smaller morphotype ∼225 μm in
length and a larger one ∼275 μm. The larger morphotype
had a chromosome number of 21 and the smaller morphotype had 14 chromosomes. The haploid chromosome number found in males of this species is n = 7. This was the first
case of polyploidy reported for rotifers, but new techniques
for staining and counting rotifer chromosomes may make
more systematic explorations possible.
Chapter | 13
243
Phylum Rotifera
Beyond morphological variation, plasticity also occurs
in sexual reproduction. Transgenerational phenotypic plasticity in the responsiveness to the mixis signal has been
described in B. calyciflorus. In some strains of this species,
females hatching from diapausing embryos do not react to
this biochemical, but, as the population develops, females
become increasingly more responsive to it. That is, sexual
reproduction is initiated, and a portion of a population
becomes sexual. Low mixis propensity continues for up to
12 generations after hatching. This suppressed responsiveness to mixis signals also has been observed in Brachionus
angularis Gosse, 1851, Rhinoglena frontalis Ehrenberg,
1853, and E. senta. Delayed mixis is regarded as an adaptive response to promote rapid asexual population growth
soon after hatching, followed by sexual reproduction at high
population densities (Serra et al., 2005). However, populations of Hexarthra sp. from temporary pools in the Chihuahuan Desert have a different mixis response (Schröder
et al., 2007). Some of the diapausing embryos that hatch in
these populations immediately go sexual, resulting in production of a new batch of diapausing embryos. Meanwhile,
the remaining newly hatched embryos reproduce asexually.
Because these small habitats may dry in a matter of days
or persist for weeks, this variation is seen as a bet-hedging
strategy: some diapausing embryos are produced immediately, while a portion of the population continues to expand
asexually.
Distribution and Population Movements
Water bodies are not uniform habitats with respect to biotic
factors such as food and predators and abiotic factors such
as dissolved oxygen concentration, light intensity, temperature, and water movements. Therefore, it is not surprising
to find that rotifers are not evenly distributed in lakes and
ponds; often there is considerable variability with respect
to their horizontal and vertical distribution. For example,
two Filinia species displayed very different vertical distribution patterns over the course of a year in a small lake in
Germany (Figure 13.18). In some meromictic lakes with
strong clinograde oxygen gradients, population maxima
may be restricted to near the oxycline. Similar patterns are
seen in the horizontal distribution of rotifers. In fact, some
rotifers are strictly littoral, being found in open waters only
as occasional migrants. Others are pelagic, but the depth at
which they are found varies with season.
Although these major distribution patterns result from
differential population growth and water currents and other
large-scale water movements, within-lake distribution patterns can be influenced to a lesser degree by the locomotory
behaviors of the animals. One commonly recognized behavior of marine and freshwater zooplankton is a daily (diel)
vertical migration (DVM) in the water column in which the
animals usually come to the surface only during the night.
During the day zooplankton avoid visual predators (fish)
FIGURE 13.18 Temporal and spatial distribution of two species of Filinia in Lake Pluβee, Germany. Solid line indicates limits of the population at
one individual per liter; dotted line indicated boundaries of higher population levels (numbers of individuals per liter). Modified with permission from
Hofmann (1982).
244
that occupy near-surface waters, but when they return to
the surface at night they can exploit the rich algal resources
present there. In rotifers, diel migrations typically are not as
dramatic as those of microcrustaceans. Thus, the population
center of rotifers usually changes only about 1–3 m over a
daily cycle. For example, a study on the zooplankton of Ross
Creek Reservoir (New Zealand) showed that Conochilopsis
causeyae (Vidrine, McLaughlin, and Willis, 1985) exhibited a nocturnal ascent with its population center ascending
from 2.1 m in the daytime to 0.9 m at night. However, two
other species did not show a nocturnal rise: Hexarthra mira
(no change) and K. cochlearis (reverse migration). In addition, ovigerous (egg-bearing) and nonovigerous females
may have different DVM patterns. Ignoring this fact can
cause serious errors in the calculations of birth rates for
field populations. Errors of nearly an order of magnitude
may occur if the population is sampled at only one depth or
at different times during the day.
Rotifers also move horizontally in aquatic systems,
with some pelagic rotifers avoiding water close to the
shore. This phenomenon, called avoidance of the shore,
was demonstrated by using a circular plexiglass arena that
permitted the researchers to assess the swimming direction of several zooplankton species over a set time interval. When the shadow produced by the natural elevation of
the shoreline was artificially altered by adding a black collar around the arena, two pelagic species, Asplanchna priodonta Gosse, 1850 and Synchaeta pectinata ­Ehrenberg,
1832, swam away from the shadow, whereas the littoral
rotifer, E. dilatata, showed no preference in its movements.
Besides the effect of the shadow cast by the shoreline, rotifer position may be explained by differences in composition of the hydrophyte community in the littoral zone.
For example, both lily pads and extensive populations of
duckweed (Lemna) provide shade under which large numbers of Conochilus hippocrepis (Schrank, 1803) may congregate (Edmondson, 1959). However, plant physiological
activities also alter the chemical properties of the water
in the immediate vicinity of the plant. For instance, polar
plants change the concentration of Ca+2 within the Prandtl
boundary (see below), and Chara ­produces a musky smell
that permeates the water.
Rotifers also are present in the psammon, the sandy habitat along the wet reaches of shorelines. While population
levels are never great in this interstitial (L., between) realm,
their diversity can be significant. For example, EsjmontKarabin (2003) found a total of 110 species in the psammon
of beaches of 18 lakes in Poland, a value representing 26%
of all species recorded from Polish waters. This community
included 22 species that occurred exclusively in the psammon, with monogononts apparently playing a much more
important role than bdelloids. Her study also reported a
tendency for higher rotifer diversity in the psammon communities of mesotrophic and eutrophic lakes as compared to
hypertrophic lakes.
SECTION | III
Protozoa to Tardigrada
Research on the abundance and distribution of rotifers
in lotic systems has lagged behind that of lentic ones. However, we do know that both rotifer diversity and abundance
are usually lower in rivers (∼5–400 individuals/l) than in
lakes, although in some instances the population density
may be quite high (ca. >6000/l). Some of the animals in
flowing waters seem to be derived from upstream lakes
or secondary channels. However, rotifers can reproduce
in both the main and secondary channels and are, for all
practical purposes, the dominant zooplankton (not counting
protists) present in certain lotic systems, such as rivers of
the Great Plains (Thorp and Mantovani, 2005). Sampling
strategies for rivers can be as complicated as those carried
out in lakes. These techniques include bank-to-bank net
tows, repeated collections at a single site, or Lagrangian
sampling. In Lagrangian sampling, the same parcel of water
is repeatedly sampled as it flows downstream.
Colonial Rotifers
Most rotifers are solitary and interact only as potential prey
or mates, but a number of species in 10 genera form intraspecific colonies or join in the formation of interspecific
colonies (Figure 13.19). Nearly all of the colonial species
are members of two monogonont families (Conochilidae and
Flosculariidae), although colony formation has been noted in
at least one bdelloid, but those colonies appear to be temporary (Wallace, 1987). All colonial species are microphagous
and many produce secretions of various sorts that link colony
mates. For example, some produce tubes from secretions that
harden (Limnias), whereas in others the secretions are gelatinous (Conochilus and Lacinularia). A few species construct
a tube out of pseudofecal pellets (e.g., Floscularia conifera
(Hudson, 1886)) or jelly and fecal pellets (Ptygura pilula
(Linnaeus, 1758)), with the resulting structures resembling
the turret of a castle. These secretions are important in the
overall structure of the colony, as they provide a substrate for
attachment of larvae or a matrix in which new members are
added to the colony (see below). Because colonial rotifers
do not reproduce by budding or the formation of specialized
zooids, colony members are not intimately connected, as are
colonial bryozoans (Chapter 16); therefore, energy resources
are not shared among colony members.
The number of individuals comprising a colony varies greatly among genera. Floscularia ringens (Linnaeus,
1758) usually builds colonies of fewer than five individuals, as do some members of the family Conochilidae
(e.g., Conochilus dossuarius Hudson, 1885). On the other
hand, some species construct colonies of intermediate size
(up to ∼35 individuals; Conochilus unicornis Rousselet,
1892), whereas large colonies (50–200 individuals) are
produced by F. conifera, S. socialis (Linnaeus, 1758), and
C. hippocrepis. A few taxa have been reported to produce
colonies of enormous size: e.g., >500; Lacinularia elliptica
Shephard, 1897.
Chapter | 13
Phylum Rotifera
FIGURE 13.19 Examples of colonial rotifers. (a) A portion of a small
sessile colony of Limnias melicerta. These animals live within a clear, firm
tube that they secrete. This species normally is solitary or forms colonies
of only a few individuals. (b) A sessile colony of Lacinularia flosculosa.
This species secretes a gelatinous matrix. (c) A tiny colony of Conochilus
(=Conochiloides) dossuarius comprising a female and her most recent offspring. As in all of the Conochilidae, this species secretes a gelatinous
matrix that is often populated by algae, bacteria, and sometimes Vorticella.
Bars = 250 μm.
So far, the mechanisms that control colony size have
not yet been explored. For example, C. unicornis usually
forms small colonies of fewer than 35 individuals; but in
four lakes with the cladoceran predator Leptodora, we
245
have collected much larger colonies from a single sample.
In those cases, the ranges of mean colony size varied from
about 82 to >200 individuals. However, our record to date
is from a population in Green Lake (Wisconsin, USA),
where the largest colonies comprised more than 400 individuals. Similar observations of >50 individuals per colony
have been made of C. unicornis populations in Canadian
Shield lakes that have the cladoceran predator Bythotrephes
longimanus. The mechanism that initiates this exuberant
colony form has not yet been elucidated, but it seems likely
that a predatory induction is responsible, as in predatorinduced spine formation in B. calyciflorus. In this case,
induction may involve an alteration in the cohesiveness of
the gelatinous matrix of the small vs. large colony forms.
Colonies form by one of three methods (Figure 13.20),
each of which produces colonies that presumably differ in
the genetic relatedness among colony mates (Table 13.1).
In allorecruitive colony formation, free-swimming juveniles produce colonies by settling on tubes of sessile
adults. Thus, colonies of allorecruitive species grow in
size (i.e., numbers of individuals per colony) by intercolonial recruitment of juveniles. The number of colonies
within the habitat (colony density) increases by juveniles
settling as solitary individuals, which are then joined by
new recruits. Because juveniles joining these colonies
may come from females belonging to another colony,
genotypic relatedness within the colony is probably relatively low. These colonies are transitory, beginning when
a recruit attaches to a previously settled adult and ending
when recruitment to an old colony ceases. Late recruits
to allorecruitive colonies may suffer because death of the
founding individual can lead to dislodgment from the substrate and subsequent sinking to the benthos. Some species of the genera Floscularia and Limnias reproduce this
way. A few species produce interspecific colonies of two
or more species, but fusion of established colonies does
not seem possible (Wallace, 1987).
In autorecruitive colony formation, the young remain
with their mother in the colony. Thus, colonies of autorecruitive species grow in size (numbers of individuals per
colony) by intracolonial recruitment of the young into the
parental colony, whereas the number of colonies within the
habitat (colony density) increases by colony fission. Because
the young joining autorecruitive colonies come from the
parent colony, genotypic relatedness within the colony is
probably high. Autorecruitive colonies are long-lived and
develop continuously throughout the season, increasing in
size as new individuals are added and decreasing only when
the colony divides. Late recruits to these colonies probably do not suffer because, although older individuals die,
the colony continues. Autorecruitive species have not been
reported to produce interspecific colonies, nor has colony
fusion been described. Species in the family Conochilidae,
as well as S. semibullata, reproduce by autorecruitive colony formation.
SECTION | III
246
Protozoa to Tardigrada
of two different sizes (ages), which is likely the result of
colony fusion (i.e., a juvenile colony joins with a previously
settled colony). Lacinularia flosculosa (Müller, 1773) and
S. socialis reproduce by geminative colony formation.
The consequences of high genetic relatedness have not
been explored, but may include increased vulnerability to
parasites, uniformity of behaviors, and decreased genetic
diversity of diapausing embryos.
Two hypotheses have been offered to explain the adaptive significance of coloniality in rotifers. One hypothesis suggests that colonial animals possess an energetic
advantage over solitary individuals of the same species.
For example, colonial F. conifera apparently live longer
and mature faster than solitary individuals. It is argued
that juxtaposition of filtering currents produced by two or
more individuals permits an increased filtering rate and/
or an enhanced filtering efficiency. Although experiments
have not supported the idea that coloniality affects filtration
rates, the dynamics of colony feeding currents suggests that
coloniality increases filtering efficiency (Wallace, 1987). A
second hypothesis argues that colonial existence can protect
individuals from certain predators. For example, large C.
hippocrepis colonies embedded in their gelatinous sheath
are less vulnerable to attack by a calanoid copepod predator
because they are too large to be engulfed whole. Also, in C.
unicornis individual rotifers retract into the refuge of the
gelatinous matrix of the colony and cannot be captured by
the predatory rotifer Asplanchna.
Sessile Rotifers
FIGURE 13.20 Three types of colony formation in rotifers: Allorecruitive
(Floscularia conifera), Autorecruitive (Conochilus), and Geminative
(Sinantherina socialis).
In geminative colony formation, all of the young born
within a span of a few hours leave the parent colony as a
free-swimming, juvenile colony. Members of this young
colony subsequently explore and attach to a new substrate
together. Thus, genotypic relatedness within a colony is
probably moderate to high, depending on its particular
history. Because size of a juvenile colony is dependent on
fecundity of its parental colony, the number of individuals
comprising a colony does not generally increase over its
lifetime. Geminative species have not been reported to produce interspecific colonies, but it is possible that they do.
We also have seen colonies that are composed of individuals
Sessile species are found in three families: Atrochidae (two
genera), Collothecidae (two genera), and Flosculariidae
(nine genera). Although they are often overlooked because
their microhabitats are generally not examined thoroughly,
these forms are actually quite common in lakes and ponds.
Occasionally they reach very high densities on plant surfaces (>6 individuals/mm2), especially in bogs and small
eutrophic ponds.
The juvenile motile stages of sessile rotifers (Figure 13.7)
are not true larvae, as all of the adult organs are present in
the young animal. However, because of the conceptual parallel with sessile marine invertebrates, and because the corona
does not appear to be completely developed in the juvenile,
the term larva is often used. Not surprisingly, the behavior of
juveniles changes dramatically once they come into contact
with a potential attachment site. These new behaviors have
been described using terms such as selection, choice, and
preference. However, use of such words is not meant to imply
cognition by the rotifer (that would be a form of teleological
thinking). These are merely convenient terms to describe this
phenomenon.
Several workers have demonstrated that juveniles can
select a particular substrate among all surfaces available
Chapter | 13
247
Phylum Rotifera
TABLE 13.1 Colony Formation in Rotifers
Colony
Formation
Category
Increase in Colony
Size (Individuals
per Colony)
Mode of Increase
of Colony Density
in the Habitat
Predicted Genotypic
Relatedness Within
Colonies
Frequency of
Interspecific
Colony Formation
Allorecruitive
(intercolonial)
Young are recruited
to established colonies
or remain solitary
Young leave the
parental colony
and may establish
a new colony
Low
Common
Floscularia (Figure
13.20); Limnias
(Figure 13.19)
Autorecruitive
(intracolonial)
Young stay within
the parental
colony
Fission: adult colonies High
fragment, producing
smaller colonies
Absent
Conochilus (Figure
13.20); Sinantherina semibullata
Geminative
Rare to absent
Cohorts of larvae
collectively establish
new colonies
Rare to absent
Lacinularia (Figure
13.19); Sinantherina socialis (Figure
13.20)
Moderate to high
for settlement. For example, juveniles of F. conifera settle
with a greater frequency on the tubes of conspecifics than
on aquatic plants, although there is substantially more plant
surface available. This propensity leads to the formation of
intraspecific colonies, each with 50 or more individuals.
During the growing season, ≥75% of the entire population
may be colonial.
Some species attach to a surface based on water chemistry. In a population of Collotheca (gracilipes) campanulata (Dobie, 1849) in a small pond filled with several
hydrophyte species, including Elodea, Lemna, Myriophyllum, and Nymphaea, most individuals were present on
Elodea canadensis (Wallace and Edmondson, 1986). In
experiments in which juvenile C. campanulata were presented with submerged Elodea, >90% of the individuals
preferred the under surface of the leaves to the upper surface (Figure 13.21). (In this case, the under surface is anatomical; it does not merely refer to the way the plant was
growing in the water, which may be sideways or upside
down due to crowding by other plants.) Attachment to the
under surface is apparently in response to the way in which
Elodea is able to alter the concentration of calcium ions in
the water immediately around the leaf, i.e., in the Prandtl
boundary layer. In water having a pH in the neutral to
alkaline range, Elodea acts as a polar plant, removing Ca+2
from beneath the leaf and releasing it above. Choice of the
under surface of the leaves of Elodea provides a superior
habitat in comparison to the upper surface. In short-term
laboratory experiments, young C. campanulata attached
to under surfaces of Elodea leaves grew significantly taller
and produced more eggs per female than those induced to
attach to the upper surfaces of the same leaves. Several
other species of sessile rotifers exhibit strong preferences
for particular substrates, but the significance of these associations has not been fully elucidated.
Examples
FIGURE 13.21 Members of the sessile species Collotheca campanulata
(=gracilipes) colonizing the undersurface of a leaf of the aquatic macrophyte Elodea canadensis. Many (>25) individuals may be seen attached to
the under surfaces of the leaf. A biofilm may be seen on the upper surface
of the distal one third of the leaf (leaf length ≈1 cm).
Substrate selection behaviors have been described for three
species: C. campanulata, P. beauchampi, and S. socialis. In
general, juveniles appear to react to potential surfaces in similar
ways. Newborns avoid settling for periods up to several hours
after hatching. Once this refractory period is past, all surfaces
are explored; however, some receive much more attention and
generate different behaviors, including some reminiscent of
male mating behavior. A juvenile will traverse these surfaces
with both its corona and foot in contact with the surface, occasionally stopping in this slightly bent position. The juvenile
may continue exploration of the surface for several minutes,
but eventually it attaches to the surface using a type of cement
from glands in the foot and then undergoes metamorphosis.
248
Immediately after attachment and metamorphosis, the
young of most sessile rotifers begin to secrete a protective
tube. Often this secretion is in the form of a clear, gelatinous
material (e.g., Collotheca and Stephanoceros; Figure 13.8),
but in some species the tube becomes obscured by colonizing microorganisms and debris (e.g., Ptygura and
Beauchampia; Figure 13.22 a). At least two sessile species augment their jelly tube with fecal pellets (Floscularia janus (Hudson, 1881) and P. pilula; Figure 13.22(b)).
Limnias melicerta Weisse, 1849 forms a cement tube that
looks like a series of transparent rings placed on top of one
another (Figure 13.22(c)). Regardless of how the tube is
formed, it probably deters some predators, but fragments
of the tubes of certain sessile rotifers may be found in snail
fecal material (e.g., Limnias).
Perhaps the most fascinating example of tube construction is found in the genus Floscularia (Figure 13.20). Species in this genus possess a small ciliated cup (modulus) on
the ventral side of the head. In some species of this genus,
the animals pass tiny particles and other small debris collected by the corona. The cilia in the modulus appear to be
in constant motion, mixing gelatinous secretions with the
particles to form small pellets (pseudofeces), either in the
shape of bullets or balls. Once a pellet is fully formed, the
rotifer places it on the top of the tube in an action resembling the movements of a bricklayer. In this way, the tube is
constantly elongated as the animal grows. Because pellets
are manufactured from particles collected from the water,
they are colored, usually light to dark brown. However,
when a heavy rainstorm temporarily suspends soil in the
water, the pellets produced by Floscularia usually turn out
to be very dark (Figure 13.23). Thus, the event of the storm
is marked as a dark ring in the tubes of all the animals alive
at that time. In his classic study, Edmondson (1945) noted
this fact, and used suspensions of powdered carmine and
SECTION | III
Protozoa to Tardigrada
carbon black to mark the tubes of F. conifera so that he
could study the dynamics of population growth in a field
population.
Another unexplored aspect of sessile rotifers is the presence of anisotropic (birefringent) crystalline structures
(ACS) that have been found in the guts of several species
(Wallace et al., 2006). Anisotropic structures possess two
to three refractive indices. Such structures may be visualized by examining a specimen set between cross polarizing filters. Under this illumination, the background is black
and the ACS bright. They also are visible under standard
bright field illumination as dark structures. When present,
ACS (ca. 10–30 μm) may take the form of small balls of
compact crystals, as in the free-swimming juvenile stages
of S. socialis, as well as in the embryos of other sessile taxa.
FIGURE 13.23 Dark pellets in the tube of Floscularia conifera. In this
instance, the animal alternately used darkly colored and lightly colored
suspended material to create its pellets. Seen here are five (left to right)
successive occurrences of dark bands. (See also Figure 13.20, top panel,
where the banding pattern is also visible.)
FIGURE 13.22 Some variation in some tube construction by rotifers. (a) Beauchampia crucigera possesses a gelatinous tube obscured by debris.
(Closed arrow = tube; open arrow = single antenna.) (b) Ptygura pilula possesses a clear gelatinous tube supplemented with fecal pellets (fp). (c) Limnias
melicerta produces a clear tube resembling a stack of rings. s = subitaneous egg. Bars = 100 μm.
Chapter | 13
Phylum Rotifera
As juvenile S. socialis age, these structures break apart into
small individual crystals that slowly disappear. However,
in the adults of the raptorial predators, Acyclus inquietus
Leidy, 1882 and Cupelopagis vorax ACS resemble compact spheres (Figure 13.24). Although present in the free-­
swimming juveniles of A. inquietus and C. vorax, these
bodies are still present in adults and enlarge as the animal
ages. In neonates, ACS may represent a source of energy
that helps the juvenile through settling and metamorphosis;
their persistence in adults may represent energy reserves
that are used during periods of starvation.
Reproduction and Life History
Reproduction
The type of reproduction varies considerably within
Rotifera. Species in class Pararotatoria (seisonids) reproduce exclusively through bisexual means, gametogenesis
occurring via classical meiosis with the production of two
polar bodies. At the other extreme, members of subclass
Bdelloidea reproduce entirely by asexual parthenogenesis
(i.e., via apomictic thelytoky). Thus, no males have been
(a)
(c)
(b)
249
observed in bdelloids, a fact that has been referred to as
an “evolutionary scandal” (Mark Welch et al., 2009). On
the other hand, species in subclass Monogononta exhibit
cyclical parthenogenesis where asexual reproduction predominates, but sexual reproduction occurs occasionally.
However, loss of capacity for sex has been documented
within laboratory populations, and males have not been
documented for all species.
Cyclical Parthenogenesis and Diapause
Cyclical parthenogenesis in monogononts (Figure 13.25)
involves an asexual (amictic) phase and a sexual (mictic)
phase. Although at certain times both asexual and sexual
reproduction occurs concurrently, in most populations only
amictic females are present. Amictic females are diploid
and produce diploid eggs termed amictic or subitaneous
eggs. These embryos develop mitotically via a single equational division and usually hatch as females within 24 hours.
Although the term summer egg has been applied to these
embryos, the term is misleading because, depending on the
species, amictic rotifers can found at any time of the year.
The switch in the reproductive mode of amictic to mictic
is initiated when an amictic female responds to a specific
stimulus (e.g., a conspecific chemical signal or an environmental cue) and begins to produce mictic daughters. These
(d)
FIGURE 13.24 Examples of anisotropic crystalline structures (ACS)
in sessile rotifers. (a) juvenile Sinantherina socialis, with ACS dispersing from its compact mass (cross polarized filters slightly offset); (b)
adult Acyclus inquietus, ACS ∼30 μm (bright field); (c) recently settled
young Cupelopagis vorax, ACS group ∼18 μm (bright field); (d) amictic
embryo of Floscularia conifer, ∼15 μm (cross polarized filters). Symbols:
arrows = ACS; int = intestine; pro = proventriculus; sto = stomach.
FIGURE 13.25 Generalized life cycle for monogonont rotifers. The life
cycle of bdelloids consists of only the parthenogenetic portion. Mixis stimulus: absent (−); present (+). Timing of mating for newborn mictic females
is critical for the outcome: early = diapausing embryos (2n), late = no fertilization, male embryos (n).
SECTION | III
250
mictic females then produce haploid eggs via meiosis that,
if unfertilized, develop into haploid males (n) and, if fertilized, develop into diapausing embryos (2n). So far, the
genetic controls of the parthenogenetic switch have not
been well examined; however, recently Hanson et al. (2013)
have shown that a cell cycle regulatory gene is important to
maintaining the typical cyclical parthenogenetic cycle.
If a male fertilizes a mictic haploid egg still early in its
development, the diploid condition is restored. However,
development of the fertilized mictic egg (zygote) is arrested
well before maturation; this results in a diapausing embryo.
These embryos possess thick walls that often are sculptured (Figure 13.26). (In some literature, the terms resting
egg, winter egg, or cyst are used to describe the diapausing embryo, but we suggest that these terms be abandoned.
Mixis is not confined to the colder months, and the term cyst
has other meanings in biology. In addition, once cell cleavage begins, a fertilized egg is properly termed an embryo.)
Diapausing embryos are very resistant to harsh environmental conditions, including desiccation, and may be dispersed
by wind (anemochory), migrating animals (zoochory), or
flowing water (hydrochory). After a period of dormancy
(a)
(b)
(c)
(d)
(e)
(f)
FIGURE 13.26 Diapausing eggs of several monogonont rotifers. (a)
Asplanchna girodi; (b) Hexarthra fennica; (c) Brachionus calyciflorus; (d)
Conochiloides natans; (e) Sinantherina socialis, (f) Kellicottia bostoniensis. (Bars = 10 μm; the wrinkled backgrounds in panels (b) and (f) are membrane filters that held the embryos during preparation.) Original scanning
electron photomicrographs courtesy of Hendrik Segers, Royal Belgium
Institute of Natural Sciences, Belgium.
Protozoa to Tardigrada
that varies among species, diapausing embryos respond to
environmental cues and hatch as diploid amictic females
(Gilbert and Schröder, 2004). The stimuli that induce hatching may include changes in light, temperature, salinity, and
oxygen concentration.
As noted above, variations exist to this typical sexual
cycle. Diapausing embryos of Hexarthra sp. from rock
pools in the Chihuahuan Desert can hatch into mictic
females, bypassing canonical mixis induction pathways
(Schröder et al., 2007). This may be adaptive for a population that lives in ephemeral habitats that may dry up within
a few days or weeks after a rainfall, facilitating rapid production of sexually produced diapausing embryos able to
withstand desiccation.
With a few notable exceptions, the stimulus for initiating sexual reproduction is poorly understood. Vitamin
E controls the shift from amictic to mictic reproduction
in some Asplanchna species, and photoperiod plays a
similar regulatory role in Notommata. A chemical signal produced by the rotifers themselves triggers mixis
in at least four species of Brachionus, two species of
Epiphanes, and R. frontalis. This process is analogous
to quorum sensing (QS) in bacteria (Snell, 2011). In B.
plicatilis, the mictic signal has been proposed to be a
protein that induces sex upon accumulating to a threshold concentration as population density increases. As
a result, rotifers are capable of auto-conditioning their
medium via secretion of the mixis induction protein(s)
(MIP). In some instances, auto-conditioned medium is
effective in inducing mixis at population densities below
about one individual per 15 ml. In fact, females can be so
sensitive to their MIP that individuals are sometimes cultured separately with bi-daily transfer to avoid accumulation of the signal to threshold concentrations. Due to
nonlinear effects, culture volume can affect the observed
density for mixis induction in laboratory experiments. In
addition to environmental factors, genetic factors play
a major role in determining the sensitivity of particular
strains to mictic stimuli. Strains that have lost the ability
to undergo mixis may still release the MIP but have lost
the ability to respond to it. At this point, the exact nature
of the MIP has not been determined.
Amictic females produce oocytes one at a time during their reproductive period, but the embryo’s fate—i.e.,
to become an amictic female or to develop into a mictic
female—depends on the concentration of the MIP to which
the mother is exposed (Gilbert, 2007). Oocytes exposed to
high levels of MIP prior to, or soon after, oviposition have a
greater chance of developing into a mictic female than those
exposed to lower concentrations. However, the probability
of amictic vs mictic progeny is not uniform over the life of
the maternal female. Offspring produced early or late in a
female’s reproductive sequence have a lower probability of
developing as mictic females.
Chapter | 13
Phylum Rotifera
Not all reproduction in monogonont rotifers follows the
pattern described above. In some populations of Asplanchna,
Conochilus, and Sinantherina, amphoteric females (individuals that produce both female and male offspring) have
been recorded (Wallace et al., 2006). Amphoteric rotifers
can produce both diploid (female) and haploid (male)
eggs, with some producing both females and diapausing
embryos, or males and diapausing embryos. The presence
of amphoteric reproduction in other rotifers has not been
fully investigated, and its significance in the life history
of those genera that exhibit it remains to be determined. A
second variation is production of eggs that resemble diapausing embryos (e.g., present of a multilayered shell) but
that were produced via parthenogenesis in the absence of
males. These eggs are termed pseudosexual eggs. In S. pectinata, two different types of parthenogenetic amictic eggs
are produced. One type resembles normal subitaneous eggs
in that they are thin shelled and develop and hatch without diapause within 1 day. Production of the second egg
type is induced by starvation. These eggs are thick-shelled
and enter an obligatory diapause after one to three cleavage
divisions. Duration of diapause in these eggs ranges from 4
to 13 days. The adaptive significance of diapausing amictic eggs seems to be to increase the ability of populations
to survive short-term food limitation. Synchaeta pectinata
also produce diapausing embryos following sexual reproduction, and these have the capacity for extended dormancy.
Diapausing Embryos in Sediments
Because of their capacity for extended dormancy, diapausing embryos theoretically can accumulate to high levels in
sediments, but the densities reported vary considerably. On
an aerial basis, densities of diapausing embryos extracted
from sediments range from 100 to 4000/cm2. Apparently,
embryos can be abundant and still viable deep into the sediments. In one study, diapausing embryos collected from at
least 21 cm deep (>40 years old) were still capable of hatching. However, reports of viability in much older sediments
have been published (Wallace et al., 2006). Of course, the
age of diapausing embryos in sediments cannot directly be
determined, but their age can be estimated from information about the sediment itself (e.g., using radiometric dating
techniques that use 210Pb and 137Cs).
Sediments from dry vernal pools in both temperate
and desert waters also will yield rotifers when rehydrated
under the proper conditions (Langley et al., 2001). Schröder
(2005) described conditions required for diapausing egg
formation, survival, and hatching, as well as possible strategies involved in these processes. No doubt the number of
diapausing embryos in basin sediments is a function of both
abiotic and biotic processes. Abiotic factors include sedimentation rate, sediment focusing (concentration of sediments in certain regions of the basin), and sediment mixing
and burial. Biotic factors include previous production levels,
251
bioturbation, hatching rates, and mortality in the sediments,
including from predation and disease.
Embryogenesis
The study of embryogenesis has not kept pace with other
aspects of rotifer reproduction. In their examination of
the bdelloid M. quadricornifera, Boschetti et al. (2005)
reported embryological studies on seven species prior to
1925 and only five more between 1956 and 1988. Most of
those studies used light microscopy on live embryos. However, confocal laser scanning microscopy has shown that
early development of M. quadricornifera is similar to that of
other rotifers: holoblastic cleavage leads to gastrulation that
occurs by epiboly. Using phallodin conjugated with rodamin, these workers visualized the mastax and trophi, noting
that these structures were formed within a mold of actin
filaments. In another study using confocal laser scanning
microscopy, Boschetti et al. (2011) identified the development stage at which the diapausing embryos of nine monogonont species became arrested. In one group, embryos
stopped cell division after about five cell divisions (mean
∼30 nuclei, with low variation in nucleus number); in the
second group cell division stopped after another division,
but the variation in nucleus number was greater (∼40–60).
Reproductive Behavior
Descriptions of mating behaviors are available for many species, including A. brightwellii, B. calyciflorus, B. ­plicatilis,
E. senta, Lecane quadridentata (Ehrenberg, 1830), Platyias
quadricornis (Ehrenberg, 1832), and Trichocerca pusilla
(Jennings, 1903). The males and females in these as in
other species show pronounced sexual dimorphism, with
males being smaller and faster swimmers (Figure 13.2).
Lacking a functional foot, males swim constantly without
attaching. Because males and females swim randomly, the
probability of male–female encounters in planktonic species can be modeled mathematically (Snell and Garman,
1986). In the colonial, sessile rotifer S. socialis, males can
copulate with several females of one colony in succession.
Females take no active role in locating a mate, but B. plicatilis females often exhibit specific reactions such as foot
flipping or accelerated swimming once a male encounter
occurs. Males, at least for some species, display a distinct
mating behavior upon encountering conspecific females.
In Brachionus, mating behavior begins when the corona
of the male squarely contacts the female. However, not all
head-on encounters result in mating; indeed, probability of
copulation in laboratory cultures generally varies from 10%
to 75% in B. plicatilis, depending on the strain. The requirement for head-on contact by the male is thought to be due
to the presence of chemoreceptors in his coronal region that
apparently respond to a species-specific glycoprotein on the
surface of the female. Mating begins with the male swimming circles around the female, skimming over the surface
252
FIGURE 13.27 Male mating behavior in brachionid rotifers. Arrows
indicate general swimming movements by male and female rotifers. (Male
not drawn to scale.)
of her lorica (Figure 13.27). During this phase, the male
maintains contact with the female with both his corona and
penis; this requires the male to remain in a slightly bent position. After several seconds of circling, the male attaches his
penis to the female, usually in the region of her corona, and
loses coronal contact. After about 1.2 minutes of copulation
in B. plicatilis, sperm transfer is completed and copulation
is terminated when the male and female break apart and
swim away. Newborn B. plicatilis males only have about
30 sperm, and they transfer two to three at each insemination.
The littoral rotifer E. senta displays an unusual mating
behavior that is unique for monogonont rotifers (Schröder,
2003). Females are mostly stationary on the substrate,
whereas males are active swimmers. If a male encounters a
conspecific female embryo, he remains near it, apparently
waiting to mate with the newly hatched female. Males discriminate between male and female eggs, and diapausing
embryos, and they exhibit a strong preference for female
embryos that are only a few hours from hatching, probably by sensing a chemical diffusing through the eggshell.
However, males have shown no difference in attending eggs
that would develop as mictic vs amictic females. Only in
the former case would a mating result in a fertilization and
production of diapausing embryos. This mating behavior is
similar to the precopulatory mate guarding of copepods, but
it lacks male monopolization of the female.
Aging
Rotifers have been used as models of aging and senescence
for a long time. Early studies on the monogonont Proales
sordida Gosse, 1886 suggested the potential for maternal
effects, in which increased age of the maternal female was
associated with a reduced overall lifespan. Concomitant
with decreased lifespan was greater variability in fecundity
SECTION | III
Protozoa to Tardigrada
and rate of development, in her progeny. This theme was
explored in a series of papers in the 1940s, which showed
that parental age also influenced longevity in the bdelloid
Philodina citrina. In these studies, orthoclones were established by isolating the first offspring of parental females,
then the first offspring of F1 females, and so forth. This
procedure yields an isogenic clone derived exclusively of
firstborn females. Orthoclones comprising the offspring of
older females also were generated (e.g., a series of the sixth
offspring). This protocol is a powerful tool for investigating effects on aging, because the only difference among
orthoclones is the age of their mothers. In P. citrina, the
mean lifespan decreased in older orthoclones, suggesting
that the phenomenon was controlled by a maternal effect
that was transmissible and cumulative. However, this effect
was reversible, because first-born females from an old
orthoclone outlived their parents. Based on these and other
studies, it was proposed that accumulation of calcium was
the maternal factor responsible. This phenomenon became
known as the Lansing Effect after the researcher who first
reported it. The rate of calcium accumulation and its importance in rotifer senescence became the reigning hypothesis
of the day. Since this early work, researchers have reported
similar results in some species but not in others.
However, a re-examination of the original data indicates
that there may not have been a direct effect on aging but,
rather, that short-lived offspring from older mothers had
increased levels of reproduction and began to reproduce
at a younger age (King, 1983). Conversely, the first reproduction of long-lived females born to young parents did
not occur until a later age. King argued that the Lansing
Effect actually could be attributed to alterations in fecundity, whereby reproduction becomes concentrated within a
few age classes in lines of short-lived rotifers. Concentrating reproduction into such an abbreviated time scale may
itself be the cause of the shortened lifespan. Until more is
known about the mechanism for these changes, King suggested that interpretations of the Lansing Effect be made
with caution.
Other experiments on aging indicate that vitamin E and
thiazolidine-4-carboxylic acid (TCA), or a combination of
chemicals can extend lifespan in both bdelloids and monogononts. Presumably, these chemicals work by quenching
free radical reactions. Two other factors that influence free
radical production are light and diet.
Lifespan is influenced by photoperiod (light:dark cycle
(L:D)), with longer photoperiods decreasing lifespan (e.g.,
short 0:24 vs long 12:12). Additionally, laboratory experiments have shown that ultraviolet radiation (UVR) shortens
lifespan significantly, with lifespan declining logarithmically as a function of dose. UVR has the potential for
damaging zooplankton in natural systems, but for rotifers
susceptibility is species-specific and also depends on water
temperature and concentration of dissolved organic carbon
Chapter | 13
Phylum Rotifera
(DOC) in the lake. For example, some species are protected from UVR by the presence of biochemicals called
mycosporine-like amino acids (MAAs) that absorb critical
wavelengths. However, MAAs are not present in all rotifers;
those that possess them do so by accumulating these biochemicals from their diet. A separate physiological response
to UVR damage is via photoenzymatic repair (PER).
Water conditions may influence light penetration into
lakes and thus the impact of UVR on zooplankton. Lakes
with significant levels of DOC, and presumably other lightabsorbing materials, attenuate UVR at shallow depths when
compared to clear water. Therefore, very low levels of DOC
in the water column have the potential to structure zooplankton species composition, favoring those species with
MAAs or the ability to undergo repair processes such as
PER, or that undergo DVM. For example, Obertegger et al.
(2008) linked the vertical distribution of rotifers in the water
column to dry weight levels of MAAs. In their study of a
clear-water oligotrophic lake Italy, they found that P. dolichoptera and Synchaeta grandis Zacharias, 1893, which
remained in the upper layer during the daytime, had ∼6 and
2 μg MAAs mg-1 dry weight, respectively.
Dietary restriction has life-extending effects in a wide
variety of animals, including rotifers. Two different modes
of dietary restriction can yield longer lifespans: reduction of
food intake and intermittent feeding. One study of dietary
restriction in rotifers indicated that restriction caused an
increase in mean and maximum lifespan, reproductive lifespan, and doubling time in most species. It seems plausible
that chronic dietary restriction is associated with changes in
reproductive allocation during starvation. Most species that
reduced their reproduction during starvation experienced
increased lifespan. In contrast, species that continued reproduction through starvation experienced decreased lifespan.
Yet, exceptions exist with some species not showing a clear
tradeoff in lifespan and fecundity.
Male lifespan has been examined far less thoroughly
than that of females, but in those species in which the male
does not feed (e.g., Brachionus manjavacas Fontaneto,
Giordani, Melone and Serra, 2007) males live only about
half as long as females (Figure 13.28). To our knowledge,
no information is available on the lifespans of males in species
that do feed (e.g., R. frontalis).
Population Dynamics
Studies of rotifer population dynamics attempt to explain
the causes of changes in population size and community
structure. By separating and quantifying the relative contributions of reproduction, mortality, and dispersal, ecologists
are able to recognize factors regulating population size and
determinants of average abundance. Because some species
are easily cultured and experimentally manipulated, rotifers
are useful models for investigating the dynamics of animal
253
FIGURE 13.28 Female and male survival of Brachionus manjavacas.
populations. Techniques also exist for investigating the
dynamics of natural rotifer populations, further broadening
their usefulness in ecological studies.
Life Tables
The most rigorous approach to studying population dynamics is life table analysis, i.e., age-specific records of all
reproduction and mortality occurring in a cohort. The general protocol for gathering life table data on rotifers is to
collect a group of newborn females, isolate them in small
volumes of culture medium (∼1–2 ml), and make daily
observations of their survival and reproduction. Maternal
females usually are transferred daily to fresh medium with a
specific food level, and offspring are counted and removed.
A table is then constructed of age, number surviving in each
age class, and number of offspring produced in each age
class. From these observations, vital population statistics
can be calculated such as mean and total lifespan, age at
first reproduction, age-specific survival (lx) and fecundity
(mx), net reproductive rate (Ro), intrinsic rate of increase
(r), generation time (T), and stable age distribution.
The detailed observations required for life table analysis usually are possible only in laboratory populations. A
number of genera have been examined in this way, including species of Asplanchna, Brachionus, and several bdelloids. In certain cases, life table analyses can be performed
on field populations, as in Edmondson’s classic study of the
colonial sessile rotifer F. conifera (Edmondson, 1945).
The major environmental factors affecting age-specific
survival and reproduction are temperature, food quantity
and quality, genetic strain, and reproductive type (either
amictic or fertilized or unfertilized mictic females).
Increased temperature shortens lifespan. For example, in
a study of B. calyciflorus by Halbach (1970), 50% of a
cohort survived to age 16 days at 15 °C, 11 days at 20 °C,
and 5 days at 25 °C (Figure 13.29). Fecundity is compressed into fewer age classes at higher temperatures.
At 15 °C, fecundity occurs at a low rate over 20 days. In
254
SECTION | III
Protozoa to Tardigrada
FIGURE 13.29 Effect of temperature on age-specific survival and
fecundity of Brachionus calyciflorus. The left Y-axis is the percentage of
the cohort surviving (open circles), and the right Y-axis is the offspring per
female per hour (closed circles). Figure redrawn from Halbach (1973).
FIGURE 13.30 Rotifer population size is determined by a variety of
factors, including food quantity and quality and levels of predation and
parasitism.
contrast, at 25 °C reproduction occurs at a high rate over
10 days. Females produce a total of 13 offspring at 15 °C,
16.6 at 20 °C, and 12.9 at 25 °C. The intrinsic rates of
increase (r) were 0.34, 0.48, and 0.82 offspring per female
per day at the respective temperatures.
Food supply obviously is an important determinant of
rotifer abundance and many studies have shown that both
the quantity and quality of the diet have an impact on population growth. However, interspecific competition (with
rotifers, cladocerans, and protists) and predation (e.g., from
cladoceran and copepods) are also important. For example,
the large predatory cladoceran Bythotrephes longimanus, a
recent invader into North American lakes, appears to have
had a significant negative impact on herbivorous and predatory cladocerans and predatory copepods in Harp Lake
(Canada). As a result, the population of the colonial rotifer C. unicornis increased significantly when released from
the pressures imposed by these crustaceans (Hovius et al.,
2007). Thus, to obtain a realistic view of rotifer growth
potential, both bottom-up and top-down processes need to
be considered (Figure 13.30).
Lake area (Ao) and food supply, as measured by annual
primary productivity (gC/m2/yr), also are important factors
in determining rotifer species richness (S). Dodson et al.
(2000) reported that there was a significant positive linear
relationship between log S and log Ao for 33 well-studied
lakes located in the northern hemisphere. They also present a regression analysis of their dataset fitted to a quadratic model, arguing that there is a unimodal relationship
between S and productivity, with the highest S occurring at
ca. 100 gC/m2/yr.
Rotifers also can detect and rapidly exploit microscale
patches (thin layers, ≤0.5 m) of food resources. In a laboratory study, individuals of B. plicatilis quickly aggregated
within thin layers of food, either Nannochloropsis oculata
or Skeletonema costatum or both, until the resource was
depleted (Ignoffo et al., 2005). From this work, we may infer
that free-swimming rotifers utilize their habitat according to
Ideal Free Distribution theory, but this is a topic that needs
greater study.
The effect of food quantity on lx and mx schedules
can be substantial. In one intensive study, Halbach and
­Halbach-Keup (1974) fed B. calyciflorus the green alga
Chlorella at densities from 0.05 to 5.0 × 106 cells/ml at 20 °C.
In that study the best survival occurred at algal concentrations
0.5 × 106 cells/ml and 1.0 × 106 cells/ml, where mean lifespan
was 9 days. Lifespan decreased at lower food levels reaching 2.5 days at 0.05 × 106 cells/ml. Short lifespans also were
recorded at food concentrations above 1.0 × 106 cells/ml,
probably the result of accumulation of algal metabolic
products that are toxic to rotifers. Fecundity also peaked at
1.0 × 106 cells/ml, with a mean of 17 offspring per female.
In contrast, at 0.05 × 106 cells/ml, lifetime fecundity was
only 0.5 offspring per female, whereas three offspring per
female were produced at 5.0 × 106 cells/ml. Similar results
have been recorded for other brachionids fed Chlorella at
densities of 106 cells/ml.
Intraspecific differences in survival among strains vary
considerably. For example, in B. plicatilis, mean lifespan
for three strains all cultured at 25 °C ranged from about
6 to 14 days. Because these strains were collected from different habitats but were acclimated to, and tested in, a common
Chapter | 13
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Phylum Rotifera
FIGURE 13.31 Life histories of Brachionus calyciflorus females grown at high (H) and low (L) food concentrations. Upper panel: amictic females producing diploid female eggs; middle panel: unfertilized mictic females producing haploid male eggs; bottom panel: fertilized females producing diploid diapausing
eggs. Numbers in parentheses are typical ranges of egg production for each female type. Percentage of life for all female types calculated based on setting the
life span of unfertilized mictic females at the high food level to 100%. Data reanalyzed from Xi et al. 2001. Internat. Rev. Hydrobiol. 86, 211–217.
environment, the observed differences must be genetic.
(Growing or culturing organisms with different genotypes
or from different habitats is typical in ecological research
and is known as the common garden technique.)
The three different female types (amictic, and unfertilized and fertilized mictic) differ markedly in their agespecific survival and fecundity. In studies on species in the
genus Brachionus, unfertilized mictic females usually lived
longer and produced more offspring than amictic females,
depending on food levels. Fertilized mictic females produced by far the fewest embryos (Figure 13.31). The greatly
reduced fecundity of fertilized females is typical of other
rotifer species and is likely due to higher energy requirements for the formation of amictic or diapausing embryos
over unfertilized mictic embryos (male eggs).
Species with different phenotypic growth forms (e.g.,
spined and unspined) can possess very different intrinsic rates
of increase under identical culture conditions. For example,
r for spined and unspined forms of K. testudo can differ significantly, but that difference depends on food concentration.
At low food levels (0.1–1 μg/ml dry algal biomass), there is
no significant difference in r for the two forms of this rotifer
(∼0.05–1). At higher food concentrations (1.2–8 μg/ml), however, the unspined form had much greater values of r than
the spined form (0.3–0.4 vs 0.15, respectively). Presumably,
similar phenomena will be found in other species in which
there are spined and unspined forms. The reader is referred to
Gilbert (2013) for a recent review of this topic.
When life-history patterns of zooplankton are compared,
rotifers have higher r than either cladocerans or copepods;
r ranges from 0.2 to 1.6, 0.2 to 0.6, and 0.1 to 0.4 offspring
per female per day, respectively. Rotifers achieve their high
population growth rates by short development times that
more than compensate for their small clutch sizes. Rotifers
also show the greatest response to increased temperatures.
The high population growth rates of rotifers may be an
adaptive response to predation, against which most rotifer
species have relatively few strong defenses. As a result of
these characteristics, rotifers are regarded as being able to
quickly exploit new conditions.
Dynamics of Field Populations
Annual cycles of natural rotifer populations have been characterized for several species, but to achieve a good appreciation of the population dynamics of rotifers long-term
studies are needed. Rotifer dynamics have been followed
for several years in Neusiedlersee, a shallow, well-mixed
Austrian lake (Figure 13.32). In this lake some species,
like K. quadrata, have relatively stable population sizes
and occupy the lake nearly continuously. In contrast, abundances of other species such as Filinia longiseta (­Ehrenberg,
1834), Hexarthra fennica (Levander, 1892), and Rhinoglena
fertoeensis (Varga, 1928) fluctuate much more, whereas
others (e.g., A. fissa) are intermittent. Fluctuation of rotifer abundance over two or three orders of magnitude during a seasonal cycle is typical of many natural populations.
Another important long-term study of zooplankton is that
of Lake Washington (Washington, USA). This work began
in 1933 and continued with intermittent sampling until
1963, when a more intensive sampling schedule was implemented. Over one 13-year period, the population level of
K. cochlearis varied by a factor of 12-fold, while another
species, C. hippocrepis, which was present only sporadically since 1933, became important (Figure 13.33).
Techniques have been developed to characterize the
dynamics of natural zooplankton populations. The intrinsic
rate of increase of a population (r) is the difference between
instantaneous birth (b) and death (d) rates, where:
r=b−d
Although more complex models can be constructed, the
value of r for a given population may be estimated from the
equation:
r = (ln Nt − ln N0 ) /t
This procedure requires two successive estimates of
population size (N0 and Nt) separated by time interval t.
The estimate of r is based on several assumptions about the
population including that it is growing exponentially with a
stable age distribution. One also can estimate the birth rate
256
SECTION | III
Protozoa to Tardigrada
FIGURE 13.32 Phenologies of several important rotifer species in Neusiedler See, Austria. Rotifer population densities are reported as individuals per
liter; note variations in the population scales. Gaps indicate missing data. Panels, from top to bottom for both columns: Total rotifer density; Rhinoglena
fertoeensis; Brachionus calyciflorus; Keratella quadrata; Keratella cochlearis; Notholca acuminata; Anuraeopsis fissa; Polyarthra vulgaris/dolichoptera; Asplanchna girodi; Synchaeta tremula/oblonga; Filinia terminalis; Filinia longiseta. The Polyarthra and Synchaeta species combinations are due to
the difficulty in diagnosing these species in preserved samples. Data courtesy of Alois Herzig, Biologische Station Neusiedlersee, Austria.
Chapter | 13
Phylum Rotifera
FIGURE 13.33 Population dynamics of two planktonic rotifers
(Keratella cochlearis and Conochilus hippocrepis) in Lake Washington
over a 13-year period. Adapted from the Lake Washington database of W.
‘Tommy’ Edmondson, as funded by the Andrew Mellon Foundation; data
courtesy of Daniel Schindler, University of Washington, WA, Seattle.
in this population by counting the number of eggs carried
per female. The finite hatching rate is:
B = E/D,
where E is the number of eggs per female and D is the
developmental rate at a specific temperature. The value E
is determined directly from samples of the population, and
values of D by observing hatching of eggs from samples
brought back to the laboratory and monitored for egg hatching. When E and D have been calculated, the intrinsic birth
rate, b, can be estimated from the following equation:
b = ln (E + 1) /D
With these data, it is possible to estimate death rate, d,
by subtraction:
d=b−r
The egg ratio technique makes it possible to predict
growth of natural rotifer populations in which reproduction
by amictic females is parthenogenetic. A few simple measurements allow ecologists to estimate important population parameters that summarize processes of birth and death
occurring in the population. Thus, this technique has proved
useful for investigating the population dynamics and secondary production of zooplankton.
As noted above, the dynamics of natural rotifer populations are affected by a number of environmental factors,
including temperature and food quantity and quality; however, exploitative and interference competition, predation,
and parasitism also are important. Temperature is a major
factor affecting fertility, mortality, and developmental rates.
In general, higher temperatures do not increase the number
of offspring produced per female; instead, they shorten birth
intervals by decreasing the development time. Lifespan also
is reduced, but the net effect of higher temperatures is an
increased population growth rate. In laboratory studies,
257
FIGURE 13.34 Synergistic effects of temperature and food density
on the intrinsic rate of population increase (rm) of Brachionus calyciflorus. From Starkweather, (1987), with kind permission of the author and
Elsevier Science and Technology Books.
higher temperatures and elevated food levels combine synergistically to increase mean rate of population increase of
B. calyciflorus (Figure 13.34). Other abiotic environmental
factors, such as oxygen concentration, light intensity, levels of pesticides, and pH, also influence rotifer population
dynamics.
Food availability is a major biotic factor regulating rotifer
population growth, and species have markedly different food
requirements for reproduction. Threshold food concentration,
the food level where population growth is zero, was determined for eight species of planktonic rotifers by ­Stemberger
and Gilbert (1985). Small species such as K. cochlearis had
lower threshold food concentrations than did larger species
such as A. priodonta or S. pectinata. The logarithm of the
threshold concentration was positively related to the logarithm of rotifer body mass, so the smallest species had the
lowest food thresholds. The food concentration required to
support 50% of the maximum population growth rate (r/2)
varied 35-fold among the eight rotifer species and also was
positively related to body mass. Small rotifer species, therefore, appear to be better adapted to food-poor environments,
because those species possess low food thresholds. Larger
species, in contrast, are better adapted to food-rich environments, where they have higher reproductive potentials.
Natural rotifer populations often are food limited. In one
study, populations of four species in two mountain ponds,
K. cochlearis, Polyarthra vulgaris Carlin, 1943, Synchaeta
oblonga Ehrenberg, 1832, and Synchaeta sp. grew faster when
provided supplemental food in enclosures. The intensity of
food limitation changed rapidly, indicating strong temporal
variation in resource availability. Although food limitation was
common, other factors such as intraspecific resource competition also strongly affected population dynamics.
Population Dynamics in Chemostats
Studies of field populations are hampered by many confounding factors, including sampling errors and variability
258
SECTION | III
Protozoa to Tardigrada
in both abiotic and biotic factors that can affect rotifer
growth. Thus, chemostats are useful tools for probing the
details of population dynamics in environments in which
abiotic factors (e.g., temperature, light, medium chemistry) and biotic factors (e.g., food quantity and quality)
can be more rigorously controlled (Boraas, 1983). Such
work has explored the conditions required for populations
to establish an equilibrium level or to cycle. In some chemostat experiments, populations have rapidly decreased or
lost their capacity for sexual reproduction. This suggests
a strong selection against the sexual cycle (see Serra and
Snell, 2009).
expression) has been used to identify genes involved in mate
recognition and thermotolerance. Comparison of sequence
variation has been used to query genes for signatures of
positive or purifying selection. Finally, next-­
generation
sequencing technologies provide an unprecedented opportunity to sequence entire genomes or transcriptomes, and
have been applied to characterize genes and gene families
expressed under conditions such as dormancy.
Genetic Variation
Initially, indirect inferences of high levels of genetic variation were derived from mating assays. These showed reproductive boundaries or inbreeding depression. However,
with the advent of electrophoretic techniques, researchers
began to study levels of allozyme variability among localities. Some of these studies revealed high levels of genetic
differentiation, whereas other studies suggested dominance of a few molecular types. Later tools for sequencing
markers (e.g., microsatellites, mitochondrial cytochrome c
oxidase genes, and rDNA) have refined the ability to characterize genetic variation within and between populations,
to discriminate among morphologically similar or identical
species (i.e., cryptic species), and to compare evolutionary
relatedness (i.e., phylogenetic analysis). Sequencing and
karyotype analyses also have increased our understanding
of the genetic structure in rotifers, discovering, for example,
the tetraploid genome of bdelloid rotifers, in contrast to the
diploid condition of monogononts.
A prominent theme in rotifer genetic analyses is the recognition of species boundaries and understanding the speciation process. Molecular tools have improved our ability
to distinguish members of cryptic species complexes, and
description of new species is ongoing. For example, the
B. plicatilis species complex is composed of three main
morphotypes (the larger B. plicatilis, intermediate-sized
Brachionus ibericus Ciros-Peréz, Gómez and Serra, 2001,
and smaller B. rotundiformis), as well as other sibling species. This group presents a classic example of morphological stasis, combined with niche partitioning thought to
facilitate species coexistence (Gómez et al., 2002). Complexes of sibling species are probably common throughout
the Rotifera, making molecular analyses critical for accurate characterization of diversity. Genetic variation among
species of bdelloids presents a particular puzzle, as these
asexual animals cannot be discriminated by the classic biological species definition of reproductive incompatibility.
Novel techniques hold promise not only for advancing
understanding of species relatedness and genetic variation,
but also for understanding functional diversification. Application of interference RNA (targeted suppression of gene
Research on food selection by rotifers began by simple
microscopic observation of rotifers feeding and by correlating the density of natural populations of rotifers to the
various types of foods available in the habitat. However, to
obtain more precise information on food selectivity, workers examined the relative number of food particles ingested
by rotifers to particle concentration in the water. This technique has been employed for both laboratory cultures and
natural populations of rotifers, and can use both natural
foods (i.e., algae, bacteria, yeast, protists) and artificial
materials (e.g., latex microspheres). To determine feeding
rates, known concentrations of food particles are provided
to rotifers, which are allowed to feed for short periods of
time. From that, the number of cells ingested per animal per
unit time (feeding rate) or the volume of water cleared of
all particles per unit time (clearance rate) can be calculated.
Natural foods also have been labeled using radioisotopes,
but loss of the radioactive label from the food can result in
significant errors when determining feeding rates. Bacteria
and algae also can be labeled by using stable isotope tracers or by using fluorescent dyes. However, even the simple
procedure of adding a few drops of food suspension to a
Petri dish with rotifers may lead to interesting observations
concerning how rotifers process food. For example, using
powdered carmine, Wallace (1987) demonstrated that adult
S. socialis do not act independently of one another when
in a colony. Instead, the corona of the animals in a large
section of the colony all face in the same direction for several minutes. Animals exhibiting this behavior form a group
called an array. The distribution of arrays within a colony
determines where the dominant water currents flow to the
colony, how the water currents flow around the colony, and
where the currents leave the colony.
Rotifers feed in ways that are directly related to their
general life history. Although most planktonic rotifers, such
as Asplanchna, Brachionus, Polyarthra, and Rhinoglena,
swim at similar rates through comparable areas of the water
column, the ways in which they encounter and capture food
can be quite different. Brachionus uses its swimming currents to sweep small food particles into the buccal region for
processing; this is often termed filter or suspension feeding.
Ecological Interactions
Foraging Behavior
Chapter | 13
259
Phylum Rotifera
Asplanchna, on the other hand, does not use its swimming
currents to gather food. Once this rotifer contacts a potential food item with its corona, it may or may not attempt
to ingest the item, based on such factors as hunger level
and the size and type of prey. Some benthic rotifers creep
along algal filaments and feed by piercing the filament and
sucking the cytoplasm from the cells; examples include
Notommata copeus Ehrenberg, 1834 and Trichocerca rattus (Müller, 1776). Sessile rotifers capture food in one of
two ways, depending on the family. Members of the family Flosculariidae (e.g., Floscularia, Ptygura, Sinantherina)
create feeding currents in a manner similar to the planktonic
suspension feeders. In contrast, all collothecid rotifers are
ambush raptors (Collotheca, Stephanoceros). Once a prey
enters the infundibular region of the corona of these rotifers,
long setae (Collotheca) or arms (Stephanoceros) fold over
the prey, capturing it much like the action of a Venus flytrap
(Figure 13.8). All members of the family Atrochidae lack
setae. For example, in C. vorax, an enlarged, umbrella-like
corona folds over the prey to capture it (Figure 13.9). In both
cases, prey is pushed through the rotifer’s mouth and into the
proventriculus where it is stored until the mastax transfers
it into the stomach. When prey are particularly abundant,
several live organisms may be seen in the proventriculus.
Occasionally, predatory rotifers such as Cupelopagis may
become so engorged that attempting to ingest another prey
item results in the loss, from the proventriculus, of one previously captured.
However, not all potential food items are consumed.
By observing individual B. calyciflorus in various densities of suspended food particles, it is possible to see that
this rotifer regulates the ingestion of food particles by three
different mechanisms. First, this species can use pseudotrochal cirri to screen certain large particles away from the
mouth. Second, particles collected by the corona may be
rejected by cilia within the buccal tube: i.e., particles that
gain entrance to the oral cavity may be removed by a reversal of the action of oral cilia. Finally, the mastax also may
actively reject particles. Members of the genera Asplanchna
and Asplanchnopus also use their trophi to remove empty
carapaces of hard-bodied prey such as other rotifers and cladocerans from their stomachs.
Although the diets of some rotifers are highly specialized, many species consume a wide variety of both plant
and animal prey and may be described as generalist feeders. However, even generalist feeders may vary in their
method of food acquisition. For example, the herbivorous
genera Brachionus and Ptygura, which process many
tiny particles in rapid succession (microphagous), have
been seen to consume small ciliates. In contrast, the main
food source of Asplanchna is often large algae; however,
despite its herbivorous diet, this genus often is considered
predatory. Perhaps a more informative term for this genus
is raptorial.
Thus, in rotifers, clear distinctions between what constitutes a predator or herbivore can be fuzzy. Therefore, it
may be more constructive to consider the ways in which
rotifers encounter and process food than what they eat. This
form of analysis is more informative about a species’ functional role in the environment. Thus, with these distinctions
in mind, it is possible to characterize the dominant trophic
type by using an index called the Guild Ratio (GR). This is
the ratio of raptorial (R) to microphagous (M) feeders. For
example, compare the trophi illustrated in Fig. 13.10 g to
13.10 i, respectively. The GR, which can be expressed either
in numbers of individuals or in biomass (e.g., R# and Rb), is
calculated as follows:
or in an alternative form,
Thus, GR′ values will range between −1 and +1, with
values <0 indicating microphagous dominance and values
>0 indicating raptorial dominance (Obertegger et al., 2011).
Given that the GR′ is determined by means of similar
information used to calculate traditional diversity indices
(e.g., S and H′), it can provide another way to follow trends
in community structure.
Molinero et al. (2005) used a similar approach in their
study of changes in the rotifer community of Lake Geneva.
This study, which was based on body size, identified a
sudden swing toward smaller-bodied rotifers beginning in
about 1987. This shift separated two regimes: an early one
(1972–1986) characterized by low temperature and high
phosphate concentrations, and a later one (1987–1998) with
higher temperature and lower phosphate levels.
Functional Role in the Ecosystem
Three groups dominate the freshwater zooplankton (protists, rotifers, and microcrustaceans), yet it is the microcrustaceans that usually receive the most attention from
researchers. Such a disparity is to be expected for two reasons, one trivial and one significant. The trivial reason is
that microcrustaceans are easier to observe, identify, and
manipulate in laboratory situations. The significant reason
is that microcrustaceans commonly account for a greater
proportion of the total zooplankton biomass than either protists or rotifers; however, this varies seasonally, and protist
and rotifer biomass can be considerably greater. Nevertheless, although protists and rotifers usually have a smaller
standing biomass than microcrustaceans, their greater numbers and high turnover rate makes them very important to
the trophic dynamics of freshwater planktonic communities, including the microbial loop (see also “Predator–Prey
Interactions”). Additionally, studies have shown that the
260
relative importance of all three taxa is often idiosyncratic
to the habitat. For example, in one study that examined the
biomass of zooplankton in two large shallow lakes, ciliates
comprised >60% of total biomass in one lake but only 6%
in the other. In the first lake, the remaining zooplankton biomass was composed of cladocerans (17%), rotifers (10%),
and copepods (9%), whereas in the second lake the pattern was cladocerans (38%), rotifers (20%), and copepods
(36%).
The importance of each species to the community also
can be visualized by ordering the species according to their
relative contribution to either community abundance or
biomass. The cladoceran Bosmina longirostris was ranked
first in density and biomass of the metazoan zooplankton in a Brazilian reservoir. However, although the rotifer
P. ­vulgaris was second in density, it was sixth in biomass
(Figure 13.35). In fact, the mean change in position of the
species based on a ranking of number of individuals to one
based on biomass was −8.2, +8.2, and +9.3, for rotifers, cladocerans, and copepods, respectively. Thus, rotifers, which
dominated the top 10 positions by abundance in this example,
did not contribute proportionally to community biomass.
Of course, knowledge of densities and biomass alone
is insufficient for understanding the functional roles of
each species in the community. Additional required information includes energy consumption and transfer through
food webs. Feeding rates of zooplankton generally are
referred to as filtration or clearance rates, and are measured
as microliters of water cleared of a certain food type per
FIGURE 13.35 Species rank–abundance and rank–biomass curves of
the summer zooplankton community (microcrustaceans and rotifers) of
Ponte Nova Reservoir, São Paulo, Brazil. Symbols: closed figures = abundance; open figures = biomass; diamonds = rotifers; circles = cladocerans;
squares = copepods (including juveniles). Numbers indicate the first four
rotifers with the highest numerical ranks. Data calculated from Table 2 of
Sendacz et al. (2006).
SECTION | III
Protozoa to Tardigrada
animal per unit of time (i.e., μl/animal/h). Usually, rotifer
clearance rates are lower than those of cladocerans and
copepods, although the rates depend heavily on food type,
temperature, and animal size. For most rotifers, clearance
rates are commonly between 1 and 10 μl/animal/h, whether
determined in the laboratory or in the field. However, a few
species can achieve levels exceeding 50 μL/animal/h.
Using estimates of clearance rates, it becomes apparent that even moderate-sized rotifers with body volumes
of about 10−3 μl process enormous amounts of water with
respect to their size: >103 times their own body volume
each hour! Ingestion rates (biomass consumed per animal
per unit time) also are very high for rotifers. An adult rotifer
may consume food resources equal to 10 times its own dry
weight per day. If their assimilation efficiencies (i.e., assimilation divided by ingestion) are between 20% and 80%, rotifers can convert a good deal of their food to animal biomass
that may be passed on to the next trophic level.
Although microcrustaceans generally have higher clearance rates than rotifers (∼10–150 and 100–800 μl/animal/h
for cladocerans and copepods, respectively), rotifers can
exert greater grazing pressure on phytoplankton than some
small cladocerans. In one study in a small eutrophic lake,
K. cochlearis populations accounted for about 80% of the
community grazing pressure on small algae during the
year. This study also showed that K. cochlearis had clearance rates about 5–13 times higher per unit biomass than
the cladoceran B. longirostris. Therefore, under certain
conditions, rotifers may be important competitors to small,
­filter-feeding microcrustaceans and are important in nutrient
recycling in aquatic systems. Furthermore, rotifers can alter
the species composition of algae in certain systems. Studies have shown that intense feeding by Brachionus rubens
Ehrenberg, 1838 can cause a shift in the dominant algal
species from Scenedesmus to the spined algae, Micractinium. Apparently this shift is based on the inability of
B. rubens to consume algae with protective spines.
Although generally not ingested, cyanobacteria are
increasingly recognized as playing an important role in
determining zooplankton species composition. Large
cladocerans are more sensitive to cyanobacteria (e.g.,
Anabaena spp.) than rotifers. The mechanism for this differential sensitivity to cyanobacteria toxicity is based on
different tendencies to ingest filamentous cyanobacteria and
different physiological tolerances to their toxins.
Thus, having the ability to reproduce rapidly, rotifers
may account for 50% or more of the zooplankton production, depending on the prevailing conditions. This production, in turn, can be an important food source for other
rotifers, Asplanchna, cyclopoid and calanoid copepods,
malacostracans (Mysis), zebra mussels, aquatic mites, insect
larvae (Chaoborus) and adults (Buenoa), and small fishes.
Abundance and species composition of rotifers often
reflect the trophic status of lakes. For example, numerous
Chapter | 13
261
Phylum Rotifera
studies have reported changes in the maximal, totalpopulation density of several orders of magnitude when
lakes were subject to intense eutrophication. Individual
species sometimes undergo dramatic population changes
during those periods. This was observed in the zooplankton of Lake Constance, where the density of Asplanchna
increased its maximum population level 280-fold over a
period of 28 year. However, in other lakes, dramatic population declines have been seen. In the years in which
in Lake Washington had elevated concentrations of dissolved phosphorus, low water transparency, and high
algal densities, K. cochlearis was abundant; however, as
these water quality parameters improved, the population
of this rotifer declined dramatically. Overall, there was
at least a 20-fold increase and then a decline during a
period of 15 years.
Studies of the interactions of rotifers with other organisms will probably continue to receive attention in the
future, especially predator–prey interactions, exploitative
and interference competition among rotifers and other herbivorous zooplankton, life history strategies, and the toxic
effects of cyanobacteria, dinoflagellates, and diatoms.
Moreover, little detailed work has been done to examine
the concept of functional complementarity in rotifer communities. This hypothesis argues that, within the constraints
of niche requirements, a decline in the population levels of
some species is offset by a rise in others, a concept termed
compensatory dynamics (Fischer et al., 2001). The idea
that the GR′ of communities can deviate widely over a season (see above) and that some rotifer communities appear
to exhibit compensatory dynamics needs to be explored in
greater detail.
extinction after 2–3 weeks (Figure 13.36). Daphnia is unaffected by the presence of rotifers. Of course, protist–rotifer
competition for bacterial food also can be important.
Population growth of certain rotifers also is inhibited by
Daphnia through interference competition. For example, in
the presence of Daphnia, K. cochlearis suffers mechanical
damage (i.e., is killed, wounded, or loses its eggs) when
swept into the branchial chamber of the daphnids; the rate
at which rotifers were killed is apparently proportional to
daphnid body length. On occasion, Keratella may be found
in the guts of Daphnia and Cypris (Figure 13.37), indicating a surprising pathway for trophic interactions (Gilbert,
2012). These microcrustaceans have the greatest impact
on Keratella populations when larger than 2 mm; similar effects have been reported for other cladocerans (e.g.,
Scapholeberis kingi). Of course, interference and exploitative competition may occur simultaneously.
Predator–Prey Interactions
Predation is another important regulatory factor in rotifer population dynamics, as rotifers are prey for several
aquatic predators including protists, other rotifers, insects,
cladocerans, copepods, and planktivorous fish. From many
studies, we know that predation affects rotifer population
dynamics both directly, by contributing to mortality, and
indirectly, as a selective force shaping rotifer morphology,
physiology, and behavior. One area that deserves additional
study is the trophic interactions of microbes, protists, and
Competition with Other Zooplankton
As noted before, rotifers, cladocerans, and copepods often
compete for limited food resources and, in general, rotifers
are relatively poor exploitative competitors because their
clearance rates are usually many times lower than those of
daphnids. In addition, rotifers also have a more limited size
range of particles that they can ingest compared to cladocerans and are less resistant to starvation. Thus, cladocerans
generally have broader food niches than rotifers in terms
of food type and size, and through direct competition may
suppress rotifer population growth. However, this outcome
may be reversed when a sufficient quantity of suspended
sediments is present in a lake.
Exploitative competition between rotifers and daphnids
is readily demonstrated when these zooplankton are grown
in single and mixed cultures. Brachionus calyciflorus and
Daphnia pulex both grow well on the alga Nannochloris
oculata in single species cultures. When both species are
present, however, Daphnia removes an increasingly larger
proportion of algal cells until the rotifers gradually starve to
FIGURE 13.36 Competition between Brachionus calyciflorus and
Daphnia pulex. Brachionus and Daphnia were grown in single species (closed symbols) and mixed-species (open symbols) batch cultures
at 20 °C, daily renewed with 5 × 106 Nannochloris cells per milliliter.
Population size (Y axis) is the number of individuals in the 80-mL culture.
Error bars (±1 SE) are visible when they exceed the size of the symbols.
Figure redrawn from Figure 1: Gilbert (1985), with kind permission of the
author and the Ecological Society of America.
SECTION | III
262
rotifers within the microbial loop (Figure 13.38) (see also
Parasitism on Rotifers below).
Most rotifers are transparent and quite small; some are
smaller than many ciliates (ca. 60–250 μm long). Although
these features benefit planktonic rotifers by reducing their
visibility to fish, a small body size renders rotifers more
vulnerable to those invertebrate predators that are tactile
feeders. Many rotifers produce a thickened integument
(lorica) and/or spines and other projections, or carry their
eggs, all of which have been shown to reduce the ability
FIGURE 13.37 Fecal pellet from Cypris pubera cultured with Keratella
tropica. Visible in this pellet are at least five loricas of K. tropica (numbers). Original photomicrograph courtesy of J.J. Gilbert, Dartmouth
College; see also Gilbert (2012).
Protozoa to Tardigrada
of predatory zooplankton to prey upon them (Wallace and
Smith, 2009). On the other hand, small size also appears to
be a deterrent to predation. In laboratory cultures, Sarma
and Nandini (2007) demonstrated that, despite its small
body size (70 μm), Anuraeopsis fissa was not consumed at
the same rate as two alternative brachionid prey by either
A. brightwellii or A. sieboldii. In fact A. sieboldii ignored
A. fissa, although it was consumed by A. brightwellii.
Rotifers have other means of reducing predatory pressures. Spines are produced in some rotifers (e.g., B. calyciflorus, Figures 13.13 and 13.39) in response to a build-up
of soluble substances released by invertebrate predators
such as Asplanchna and several genera of copepods Epischura, Mesocyclops, and Tropocyclops. This phenomenon is
another type of chemical signaling, except that the communication is between predators and prey rather than among
conspecifics as in quorum sensing.
Polymorphic spine production has been observed in
B. calyciflorus, Brachionus urceolaris Müller, 1773,
F. longiseta, K. cochlearis, Keratella slacki Bērziņš, 1963,
and K. testudo. The importance of spined morphotypes is a
significant reduction in capture and ingestion by invertebrate
predators by making the rotifer more difficult to manipulate and swallow. The presence of spines on B. calyciflorus
is a good example of the phenomenon (Figure 13.13) that
works as follows. When disturbed by a potential predator, B. calyciflorus will retract its corona and, by doing so,
increases the hydrostatic pressure within its pseudocoelom.
(a)
(c)
(b)
(d)
FIGURE 13.38 Examples of rotifers in the microbial loop. (a) Heliozoan with ingested Lecane. (b) Ciliate (Frontonia) with ingested Lecane.
Photomicrograph courtesy of John Maccagno. (c) Asplanchna with two Keratella within its gut. (d) Several individuals of an unidentified microbe within
the lorica of a dead Euchlanis. This culture crashed within three days; all of the dead rotifers were infested like this one. Key: r = rotifers; bar = 100 μm.
Chapter | 13
Phylum Rotifera
The elevated pressure causes the posteriolateral spines to
swing forward and outward (Figure 13.39), making it more
difficult for a predator to manipulate. For Asplanchna,
this change is sufficient to prevent ingestion after capture.
After a period of time, during which Asplanchna attempts
to swallow B. calyciflorus (usually >60 s), the predator will
release (reject) the prey, which then swims away unharmed.
In this example, Asplanchna releases a biochemical cue (an
allelochemic, called a kairomone) that initiates a developmental change in subsequent generations of B. calyciflorus
(spine production), reducing the effect of predation. Some
forms of K. cochlearis also possess posterior spines that
make them much more likely to be rejected after capture
by Asplanchna girodi than unspined forms. Unfortunately,
the cost of spine production in rotifers, either in terms of
developmental costs of producing the spines or increased
energetic demands on swimming by increased mass, has
not been fully resolved (Gilbert, 2013). Sometimes when
Asplanchna or a related species (Asplanchnopus) swallows
a spined form, the spines become lodged in the gut of the
predator and may even puncture the delicate tissues. Presumably, both predator and prey die when that happens.
Similarly transparent mucus sheaths produced by some
planktonic, colonial rotifers (e.g., Conochilus, F
­igure
13.19(c), and Lacinularia) deter predation; the sheaths
make the effective size of the prey too large for the invertebrate predator (e.g., Asplanchna and predatory copepods)
without making them more visible to fish. The tubes and
sheaths of sessile rotifers also work as refugia (Figures
13.8(a),(d) and 13.19(a)).
Some rotifers escape predators by making rapid jumps
using a variety of appendages: setous arm-like appendages
(Hexarthra, Figure 13.14), paddles (Polyarthra, Figure
13.40), and long setae (Filinia, Figure 13.41). However,
apparently Filinia also can use its long setae as foils to ward
off predators and not just as lever-arms to initiate a jump.
FIGURE 13.39 Extension of the posterolateral spines in Brachionus
calyciflorus. Here the spines are extended due to preservation in formalin.
In life retraction of the corona causes an increase in the pressure within the
pseudocoelom, which results in an outward flexing of the spines.
263
Some rotifers assume a passive posture, displaying what has
become known as the dead-man response, rather than fleeing when predators attack (Asplanchna, Brachionus, Keratella, Sinantherina, and Synchaeta). This simple behavior is
FIGURE 13.40 Polyarthra. This rotifer possesses long paddle-like
appendages that are used in making rapid jumps to escape predators. Photomicrograph courtesy of Martin V. Sørensen, University of
Copenhagen.
FIGURE 13.41 Filinia. This rotifer possesses long spines that it uses as
foils to rapidly fend off predators. Photomicrograph courtesy of Martin V.
Sørensen, University of Copenhagen.
264
SECTION | III
Protozoa to Tardigrada
members of the Scenedesmaceae (Chlorophyta). For
example, Scenedesmus obliquus exhibited a logistic dose–
response to dilutions of test medium in which B. calyciflorus
was incubated, thus indicating the presence of grazingreleased biochemicals, termed infochemicals (Verschoor
et al., 2004). Algal defenses such as these can influence the
long-term stability of both rotifer and algal populations and,
in doing so, steady the population fluctuations often seen
in bi- and tritrophic food chains. Rapid evolution also has
been documented in long-term chemostats in which rotifers
were fed algae from a stock composed of multiple clones. In
the presence of rotifer grazing, there is strong selection for
algal genotypes that are more digestion resistant; however,
this defense comes at a cost of slower population growth
(Yoshida et al., 2004).
Parasitism on Rotifers
FIGURE 13.42 A few individuals of a colony of Sinantherina socialis.
Here all the animals have retracted their corona, a behavior that exposes
the warts on their anteroventral surface.
merely a retraction of the corona into the body and passive
sinking. Contraction of the corona stops the animal from
swimming, which eliminates the vibrations it produces that
may be detected by the predator. In addition this behavior
may make the rotifer more difficult to grasp in its turgid
state. In Sinantherina spinosa (Thorpe, 1893), this passive
posture exposes a group of small spines on its anteroventral
body surface that may function in defense against planktivorous fish.
To date, only one species of rotifer, S. socialis, has been
shown to be unpalatable to small zooplanktivorous fishes
(Felix et al., 1995) and certain invertebrates (Walsh et al.,
2006). While neither the nature nor the location of the
unpalatability factor(s) is known, this colonial species probably possesses a chemical that is held in gland-like structures, called warts, located at the anterior end of the animals
(Figure 13.42).
Defenses against predators such as spines, mucus
sheaths, thickened loricas, and escape movements are energetically demanding. Some species, such as Synchaeta pectinata, are not well defended against predators, but have
evolved very high maximal population growth rates that
offset mortality from predation. Avoiding potential predators in space and/or in time is another simple yet effective
defense mechanism against predators. Some rotifers occupy
the habitat at a different time of year from that of an important predator, migrate vertically or horizontally in the habitat, or live in zones with low oxygen concentration, thereby
evading predatory pressures altogether.
Some algae initiate colony formation as an antipredator defense when rotifers feed on them. This occurs in
The importance of parasites in controlling population density
in rotifers has not been examined thoroughly, although a few
studies have correlated parasitic infection with a decrease in
population density of planktonic species. In certain cases,
parasites apparently caused the demise of an entire population in a lake within a few days. At least one virus causes high
mortality in B. plicatilis aquaculture systems. The s­ porozoan
parasite Microsporidium (Plistophora) frequently infects
planktonic rotifers possessing thin loricas, such as members of the genera Asplanchna, Brachionus, C
­ onochilus,
­Epiphanes, Polyarthra, and Synchaeta. Water temperature
seems to be an important mediating factor in the spread of
the parasite, as infection rates drop off at water temperatures
below 20 °C. In infected rotifers, the pseudocoelom of the
animal becomes nearly filled with cysts (Figure 13.43).
Several workers have described endoparasitic fungi that
attack soil rotifers of the genera Adineta and P
­ hilodina.
Some fungi form peg-like adhesive appendages on both
small conidia (spores, ca. 30 μm long) and long vegetative hyphae. Once the adhesive pegs attach to a rotifer,
they germinate and rapidly colonize the pseudocoelom
(Figure 13.44). Other fungi produce spores that initiate
parasitic attack when ingested. Another avenue of infection occurs via hypodermic injection of a vegetative cell
into the host. Once inside the host, these fungal cells grow
into ­assimilative hyphae, producing more infective cells
either inside or outside the rotifer. However, bdelloids
apparently free themselves from fungal parasites through
anhydrobiosis (Wilson and Sherman 2010, 2013).
Rotifers also can ingest the oocysts of parasitic protists
important to human health such as Cryptosporidium and
Giardia. However, it is not known whether they can significantly reduce the numbers of these parasites in natural conditions. Rotifers also act as vectors of disease agents such
as white spot syndrome virus, which have been reported to
attack shrimp in aquaculture systems.
Chapter | 13
265
Phylum Rotifera
FIGURE 13.45 Phoretic association between Brachionus and Daphnia.
Photomicrograph courtesy of Elizabeth J. Walsh, University of Texas at
El Paso.
FIGURE 13.43 Photomicrograph of Brachionus sp. infected with a sporozoan parasite within its body cavity.
termed phoresis (G., to carry), to either the daphnid or to
the rotifer have not been completely explored.
COLLECTING, CULTURING, AND
PREPARATION FOR IDENTIFICATION
Collecting
FIGURE 13.44 An example of a fungal parasite of rotifers. Four rotifers trapped by adhesive pegs on the vegetative hyphae of Cephaliophora.
Bar = 100 μm. (Redrawn with permission of George Barron and the
Canadian Journal of Botany 61(5):1345–1348.)
Rotifers as Parasites
A few rotifers have been described as being parasitic on
algae, sponges, other rotifers, freshwater oligochaetes, snail
eggs, crustaceans, and fishes. However, we really know
too little about these associations to classify all of them
as parasitic. Members of the genera Brachionus, Limnias,
Pleurotrocha, Proales, and Ptygura have been known to
make temporary attachments to either invertebrates or vertebrates. In fact, it is not uncommon to find the carapace of
Daphnia colonized by numerous individuals of B. rubens
­(Figure 13.45). The consequences of this phenomenon,
Collecting rotifers does not require complex or expensive equipment (Wallace et al., 2006; Wallace and Smith,
2009). One can almost always collect several species of
planktonic rotifers by towing a fine-mesh (25–50 μm) net
through any body of water. It is important to note that nets
with larger mesh sizes (≥63 μm) tend to miss small-bodied
forms (Chick et al., 2010). Productive lakes and ponds usually provide especially good sampling sites, but because
fine-mesh nets can clog easily in these habitats, the water
may need to be prefiltered through a net with a larger mesh
size. More elaborate equipment such as closing nets and the
Clarke-Bumpus sampler work well, but they are not necessary unless required by a specific sampling protocol. Water
collected by discrete sampling devices (e.g., Van Dorn sampler, Kemmerer bottle, plankton trap, submersible pump) is
then filtered. In weedy areas, a dip net or flexible collecting
tube, called a water core, are very useful.
Another simple method to collect rotifers is to submerge
a 3- to 4-l (1-gallon) glass jar in a weedy region and to
arrange loosely a few aquatic plants in it before retrieval.
In the laboratory, place the jar near a subdued light source
such as a north-facing window or a low-intensity lamp.
Rotifers that swim to the surface on the lighted side may
be removed using a transfer pipette. Certain aquatic plants
such as Elodea, Myriophyllum, and filamentous algae are
SECTION | III
266
good substrata to examine for the presence of sessile rotifers, but Utricularia usually provides the richest diversity
(Edmondson, 1959; Wallace, 1980). Plants with highly dissected leaves may be examined in small dishes using a dissecting microscope. Broad-leaved plants must be cut into
strips and examined on edge.
The upper few centimeters of moist sand taken just
above the water line along a lake or a marine shoreline or
the hyporheic interstitial zone of a streambed usually provides several species of rotifers. Unfortunately, very few
studies have been conducted on rotifers from the psammon,
in part because of the difficulty in separating the organisms
from the sand. Recent studies have revealed a remarkable
diversity (35–85% of the fauna) and abundance (up to 105
individuals/l) of rotifers in this habitat. Under certain conditions, rotifers may be found at depths of up to 60 cm into
the interstitial.
Sediments collected from the bottom using a dredge,
coring apparatus, or suction device usually provide several species. The upper few centimeters of sediments from
a core normally contain diapausing embryos that can be
induced to hatch within a few days when several milliliters
of sediment are incubated at ambient spring or summer temperatures. Dry sediment from desert rock pools also provide
material when rehydrated in culture fluid and incubated for
several days.
Do not overlook laboratory aquaria as potential sources
of material. We have found some unusual species in aquaria
that had remained almost unattended for months. One might
try adding a small amount of sediments from several sites as
a way of adding variety to the rotifer community within the
aquarium. Sessile rotifers may be present if the aquarium
contains aquatic plants recently collected from the field.
However, if you attempt to keep sessile forms, be sure
to remove snails from the aquarium. Aquatic plants from
hobby shops may bring new rotifer taxa to your aquarium,
but the plants themselves may be alien species. Exercise
care so that they are not released into the environment. Populations of rotifers may be maintained or even increased by
adding a small amount of food once or twice a week (see
Culturing), but keep the aquarium aerated. Aquarium filters
that use fibrous materials to remove suspended materials
may reduce the rotifer population.
Once the collection has been made, it should be examined alive as soon as possible. It is generally a good idea to
place live samples in jars over ice for the return trip to the
laboratory, although we have found that a few species suffer when cooled (i.e., collections made from warm waters).
Anesthetizing rotifers in the field has proved to be difficult
(see below).
Three preservatives are commonly used to preserve
rotifers: formalin and Lugol’s iodine (I2KI) at concentrations of 5% or less, and ethanol at about 30–50%. (Higher
concentrations of ethanol are used in preservation for DNA
Protozoa to Tardigrada
analysis.) Lugol’s has two advantages over formalin. It is
less toxic, and it stains the specimens slightly, which makes
the animals more visible during sorting. Unfortunately,
fixation deforms the specimens, making them difficult, if
not impossible, to identify. On the other hand, in the genus
Lecane, fixation with formalin is advisable so that one can
study the morphology of the lorica. Dyes such as Rose
Bengal are commonly used to stain preserved specimens.
However, preservation sometimes causes rotifers to stick
together and to other zooplankton. This preservation artifact
has been misinterpreted as a behavior, as is seen in case of
live B. rubens, which attaches to free-swimming Daphnia
(Figure 12.45).
Culturing
Laboratory Culture
Many species of rotifers have been cultured for research,
most notably in the genus Brachionus, both freshwater
(B. calyciflorus and B. rubens) and saline (B. plicatilis).
Because of their extensive use, these species have been
referred to as the white mice of the rotifer world. The culture
systems can be quite simple, using only small vessels such
as depression slides, watch glasses, plastic tissue culture
plates (Figure 13.46(a)–(b)), and small beakers or flasks
(Figure 13.46(c)). However, larger systems are often used
(Figure 13.46(d)), and, in aquaculture, very large systems
are employed (Figure 13.46(e)–(f)). Wallace et al. (2006)
provides a summary of procedures for culturing rotifers for
research or aquaculture settings. Carlson (2000) provides
directions for construction of a simple system for the hobbyist, which can be scaled up for research or large-scale culture (see Lawrence et al., 2012). Culture vessels that have
had contact with formalin should not be used, as a residue
of this toxin is thought to remain attached to both glass and
plastic.
Some species easily adapt to artificial conditions, attaining densities of >105 individuals per liter in a few weeks,
whereas others seem impossible to keep even for short periods. Most cultures need regular maintenance a few times
per week (e.g., changing the medium, feeding, cleaning
the vessels). However, some species require little care. For
example, some bdelloid species (e.g., Habrotrocha rosa,
which is found in pitcher plant traps) are easily cultured in
dilute suspensions of powdered baby food or crushed dried
pet food. In this case, decomposition of the food source provides bacteria for the rotifers. In a similar manner, the techniques used to culture protists (e.g., making extracts and
infusions of various grains, hay, manure, soil) have been
adopted for the culture of some rotifer species with great
success. Such cultures may be ignored for days and perhaps weeks at a time without loss of the culture. Nevertheless, most researchers grow the food needed to culture their
Chapter | 13
267
Phylum Rotifera
(a)
(b)
(c)
(d)
(e)
(f)
FIGURE 13.46 Different scales in culturing rotifers. Small-scale laboratory cultures: (a) Plastic tissue culture plates are common for small studies.
Here, a 12-well plate (effective volume ≤5 ml) is shown, but 48-well plates (∼1 ml) are used to culture individual animals. (b) Plastic tissue culture flasks
are used to achieve slightly larger populations (ca. 250 ml). Medium-scale laboratory cultures. (c) In this two-stage culture system, algae are grown in
plastic bags on one shelf (numbers indicate relative age; oldest = 1), which is slowly supplied to the small columns (∼575 ml) on the lower shelf by gravity feed or a pump. Larger laboratory systems: (d) A single-stage culture system in which algae are grown in 250-l plastic columns and then a starter
population of rotifers is added. Aquaculture scales: (e) High-density mass culture (>1 m3) achieve up to 4000 individuals/l (Nagasaki Prefectural Institute
of Fisheries). (f) Mass culture pools (50 m3) achieve up to 400 individuals/l (Japan Sea Farming Association). Photographs (e) and (f) courtesy of Atsushi
Hagiwara, Nagasaki University.
target species separately and feed it to the culture at regular intervals, usually 2–3 times per week. Although more
costly, commercial products such as Roti-Rich® (Florida
Aqua Farms, Inc.) and Sparkle® and related products (INVE
Aquaculture®) give excellent results, especially where large
numbers of rotifers are required. However, do not overlook
the prospect of large populations suddenly arising in fish
tanks. One of us (R.L.W.) followed the population dynamics of the sessile rotifer C. vorax (ca. 200–1100 μm) on the
sides of a 115 L (50-gal.) aquarium for nearly 7 years; during that time, the density varied from <0.1 to ∼20 individuals per square centimeter.
Most rotifer cultures are maintained as xenic systems
without much of a problem. In fact, sometimes rotifers are
found contaminating cultures of protists, microcrustaceans,
etc., that have been provided by commercial biological
supply companies. Rotifers also have been maintained
under axenic or monoxenic culture conditions, but these
require much more effort. Sophisticated culture techniques
using single- and two-stage chemostats have been discussed
elsewhere in this chapter.
Aquaculture
Many small fish are well adapted to locate, pursue, capture, and ingest microzooplankton, including rotifers.
These organisms are easy prey because they swim slowly
and frequently lack sufficient predator defenses. Aquaculturists have exploited this important relationship between
planktivorous fish and rotifer prey in intensive aquaculture systems in both freshwater and marine systems
­(Figure 13.46(e)–(f)). This field has developed into a major
268
technical discipline in several countries including China,
India, Israel, Japan, and several countries in Europe. Most
of this work has the practical goal of determining the correct biotic and abiotic factors necessary to maintain mass
cultures of rotifers. The rotifers are then provided as the first
food for larval stages of crustaceans and fishes. In general,
rotifers are highly nutritious, and their biochemical composition can be furthered improved by specialized diets.
Most systems for mass culturing of rotifers are simple batch
cultures capable of producing kilogram quantities of rotifer
biomass each day.
SECTION | III
Protozoa to Tardigrada
crushing or distorting the specimen with the cover glass.
Both objectives may be accomplished with a compression
microscope slide. If a compressor is unavailable, then tiny
pieces of broken cover glass (not recommended) or little
clay corner supports (recommended) work well to elevate
slightly the cover glass. If the clay supports are too high,
a slight pressure from a pencil on each corner will reduce
the height of the cover glass to the desired level. With some
practice, one can trap a planktonic rotifer sufficiently to prevent swimming without undue constriction. Sessile rotifers
are handled more easily. Plant material with attached rotifers can be trimmed with iridectomy scissors to a size suitable for placement on a microscope slide. The animal will
Preparation for Identification
remain in place without the need of compression as long as
Unless special precautions are taken before fixation, illoricate the plant material is large enough to act as an anchor. Clay
rotifers (especially bdelloids) will contract into a completely supports can help level the cover glass as necessary.
unidentifiable lump, making identification difficult, if not
Methylcellulose or other viscous agents and fibrous
impossible. Such specimens have lost all value for taxonomic material, such as glass wool or shredded filter paper, also
purposes, although the trophi still may be useful. Sugar–­ may be used to impede swimming species. Unfortunately,
formalin solutions, which are used to prevent osmotic shock methylcellulose interferes with ciliary function, and fibers
in cladocerans (e.g., Daphnia), are not helpful in preventing reduce observation to a game of hide-and-seek. Any lightcontraction in rotifers. Whenever possible, live specimens ing conditions may be used, as long as you are careful not to
should be examined first, then fixed to determine the effect overheat the specimens. Strobe lighting provides a marvelof a particular fixative on body shape. In some forms, identi- ous view of ciliary movements, and dark field illumination
fied based on the shape of their lorica (e.g., Lecane ­species), is often spectacular!
formalin fixation is required (see above).
The final identification of many rotifer species requires
Anesthetics ranging from simple carbonated water to examination of the trophi, which, in certain species, may be
chemicals that are controlled substances have been used done by compressing an intact animal (e.g., Asplanchna).
to anesthetize and sometimes to kill rotifers, but none of However, one may extract trophi from surrounding soft tisthese are universally effective (Edmondson, 1959; Wallace sues using a small volume of bleach (sodium hypochlorite).
et al., 2006). Adding minute amounts of powdered MS-222 Some descriptions of this technique use a depression slide,
(Tricaine), a fish anesthetic, over the course of 30–45 min but that requires that the trophi be moved to a regular slide
to a small drop of water works well for some species. Some after the hydrolysis is complete; this is a very difficult task.
researchers use this technique, but instead substitute minute We recommend using a regular microscope slide from the
drops of formalin over longer periods of time. This takes start, but without the cover glass. Because the trophi are
extreme patience and still may not work. Carbonated water liberated rather quickly when the bleach comes into contact
(club soda) may be used to anesthetize rotifers, as well as with the rotifer, it is necessary to find the animal rapidly;
microcrustaceans. Apparently the elevated CO2 causes otherwise, it becomes necessary to scan the entire slide
asphyxiation, which may be only temporary. Unfortunately, for the small trophi. Be aware that bubbles may form and
adding carbonated water to the edge of a cover glass results in obscure your view when bleach comes into contact with
production of annoying bubbles that can hinder one’s view. some biological materials. In all work with trophi, it is
An alternative approach that bypasses the need for anestheti- important to remember that these structures are very small
zation is to use the hot-water fixation technique (Edmondson, (<50 μm) and that they are three-dimensional objects with a
1959). This technique generally gives good results with a particular spatial arrangement among all seven pieces.
number of species, once it has been mastered. However, even
DNA barcoding has been applied to rotifers with some
when great care is taken to anesthetize a rotifer, the striking success, but it has not yet been widely adopted and has a
beauty of the living animal is lost by any fixation and mount- long way to go before it will be practical for identification
ing procedure. Techniques for anesthetization, preservation, of species. In early studies, the mitochondrial gene for suband mounting of rotifers for examination using light micros- unit 1 of cytochrome oxidase (cox1, also mtCOI) was used
copy and transmission and scanning electron microscopy almost exclusively and worked well on a variety of bdelloids
have been reviewed by Wallace et al. (2006), and Jersabek and monogononts. Of course, DNA extractions can be done
et al. (2010) provides a detailed protocol.
successfully only with animals that have been preserved in
Observations of live rotifers are not always easily ethyl alcohol (95%). We believe that an experienced taxonaccomplished. The goal is to retard their movements without omist should be consulted to confirm species identifications
Chapter | 13
Phylum Rotifera
early in genetic work, and certainly well before sequences
are deposited in GenBank®.
REFERENCES
ASTME 1440–91. ASTM, 2012. ASTM. Standard guide for acute
­toxicity test with the rotifer Brachionus. Am. Soc. Test. Mater. (West
­Conshohocken, Pennsylvania).
Błędzki, L.A., Ellison, A.M., 2002. Nutrient regeneration by rotifers
in New England (US) bogs. Verhandlungen Int. Ver. Limnol. 28,
1328–1331.
Boraas, M.E., 1983. Population dynamics of food-limited rotifers in a twostage chemostat culture. Limnol. Oceanogr. 28, 546–563.
Boschetti, C., Leasi, F., Ricci, C., 2011. Developmental stages in diapausing eggs: an investigation across monogononts rotifer species. Hydrobiologia 662, 149–155.
Boschetti, C., Ricci, C., Sotgia, C., Fascio, U., 2005. The development
of a bdelloid egg: a contribution after 100 years. Hydrobiologia 546,
323–331.
Carlson, S., 2000. How to rear a plankton menagerie. Sci. Am. 283, 84–85.
Chick, J.H., Levchuk, A.P., Medley, K.A., Havel, J.H., 2010. Underestimation of rotifer abundance a much greater problem than previously
appreciated. Limnol. Oceanogr. Methods 8, 79–87.
Clément, P., Wurdak, E., 1991. (Chapter 6) Rotifera. In: Harrison, F.W.,
Ruppert, E.E. (Eds.), Microscopic anatomy of invertebrates. Aschelminthes, vol. 4. Wiley-Liss, Inc., New York, NY, pp. 219–297.
De Meester, L., Gómez, A., Okamura, B., Schwenk, K., 2002. The monopolization hypothesis and the dispersal–gene flow paradox in aquatic
organisms. Acta Oecol 23, 121–135.
Denekamp, N.Y., Reinhardt, R., Kube, M., Lubzens, E., 2010. Late
embryogenesis abundant (LEA) proteins in nondesiccated, encysted,
and diapausing embryos of rotifers. Biol. Reprod. 82, 714–724.
Dodson, S.I., Arnott, S.E., Cottingham, K.L., 2000. The relationship in
lake communities between primary productivity and species richness.
Ecology 81, 2662–2679.
Dolan, J.R., Gallegos, C.C., 1992. Trophic role of planktonic rotifers in
the Rhode River Estuary, spring - summer 1991. Mar. Ecol. Progess
Ser. 85, 187–199.
Edmondson, W.T., 1945. Ecological studies of sessile Rotatoria, Part II.
Dynamics of populations and social structure. Ecol. Monogr. 15,
141–172.
Edmondson, W.T., 1959. (Chapter 18) Rotifera. In: Edmondson, W.T.
(Ed.), Freshwater biology, second ed. John Wiley & Sons, Inc., New
York, NY, pp. 420–494.
Epp, R.W., Lewis Jr, W.M., 1980. Metabolic uniformity over the environmental temperature range in Brachionus plicatilis (Rotifera). Hydrobiologia 73, 145–147.
Epp, R.W., Lewis Jr, W.M., 1984. Cost and speed of locomotion for rotifers. Oecologia 61, 289–292.
Epp, R.W., Winston, P.W., 1978. The effects of salinity and pH on the
activity and oxygen consumption of Brachionus plicatilis (Rotatoria).
Comp. Biochem. Physiol. 59A, 9–12.
Esjmont-Karabin, J., 2003. Rotifera of lake psammon: community structure versus trophic state of lake waters. Pol. J. Ecol. 51, 5–35.
Eyres, I., Frangedakis, E., Fontaneto, D., Herniou, E.A., Boschetti, C.,
Carr, A., Micklem, G., Tunnacliffe, A., Barraclough, T.G., 2012. Multiple functionally divergent and conserved copies of alpha tubulin in
bdelloid rotifers. BMC Evol. Biol. 12, 148.
269
Feike, M., Heerkloss, R., 2009. Does Eurytemora affinis (Copepoda) control the population growth of Keratella cochlearis (Rotifera) in the
brackish water Darß–Zingst Lagoon (southern Baltic Sea)? J. Plankton Res. 31, 571–576.
Felix, A., Stevens, M.E., Wallace, R.L., 1995. Unpalatability of a colonial
rotifer, Sinantherina socialis, to small zooplanktivorous fishes. Invertebr. Biol. 114, 139–144.
Fischer, J.M., Frost, T.M., Ives, A.R., 2001. Compensatory dynamics in
zooplankton community responses to acidification: measurement and
mechanisms. Ecol. Appl. 11, 1060–1072.
Fontaneto, D., Barbosa, A.M., Segers, H., Pautasso, M., 2012. The ‘rotiferologist’ effect and other global correlates of species richness in monogonont rotifers. Ecography 35, 174–182.
Frost, T.M., Fischer, J.M., Klug, J.L., Arnott, S.E., Montz, P.K., 2006. Trajectories of zooplankton recovery in the Little Rock Lake whole-lake
acidification experiment. Ecol. Appl. 16, 353–367.
Gilbert, J.J., 1967. Asplanchna and postero-lateral spine production in
Brachionus calyciflorus. Arch. für Hydrobiol. 64, 1–62.
Gilbert, J.J., 1980. Female polymorphisms and sexual reproduction in
the rotifer Asplanchna: evolution of their relationship and control by
dietary tocopherol. Am. Nat. 116, 409–431.
Gilbert, J.J., 1985. Competition between rotifers and Daphnia. Ecology
66, 1943–1950.
Gilbert, J.J., 2007. Induction of mictic females in the rotifer Brachionus:
oocytes of amictic females respond individually to population-density
signal only during oogenesis shortly before oviposition. Freshwater
Biol. 52, 1417–1426.
Gilbert, J.J., 2012. Effects of an ostracod (Cypris pubera) on the rotifer
Keratella tropica: predation and reduced spine development. Int. Rev.
Hydrobiol. 97, 445–453.
Gilbert, J.J., 2013. The cost of predator-induced morphological defense
in rotifers: experimental studies and synthesis. J. Plankton Res. 35,
461–472.
Gilbert, J.J., Schröder, T., 2004. Rotifers from diapausing, fertilized eggs:
unique features and emergence. Limnol. Oceanogr. 49, 1341–1354.
Gilbert, J.J., Williamson, C.E., 1983. Sexual dimorphism in zooplankton (Copepoda, Cladocera, and Rotifera). Annu. Rev. Ecol. Syst.
14, 1–33.
Gómez, A., Serra, M., Carvalho, G.R., Lunt, D.H., 2002. Speciation in
ancient cryptic species complexes: evidence from the molecular phylogeny of Brachionus plicatilis (Rotifera). Evolution 56, 1431–1444.
Halbach, U., 1970. Die Ursachen der Temporal Variation von Brachionus
calyciflorus Pallas (Rotatoria). Oecologia 4, 262–318.
Halbach, U., 1973. Life table data and population dynamics of the rotifer
Brachionus calyciflorus pallas as influenced by periodically oscillating temperature. In: Wieser, W. (Ed.), Effects of Temperature on Ectothermic Organisms. Springer-Verlag, Berlin, pp. 217–228.
Halbach, U., Halbach-Keup, G., 1974. Quantitative Beiehungen zwischen
Phytoplankton und der Populationdynamik des Rotators Brachionus
calyciflorus Pallas. Befunde aus Laboratoriums-experimenten und
Freilanduntersuchungen. Arch. für Hydrobiol. 73, 273–309.
Hanson, S.J., Schurko, A.M., Hecox-Lea, B., Mark Welch, D.B., Stelzer,
C.-P., Logsdon Jr, J.M., 2013. Inventory and phylogenetic analysis of
meiotic genes in monogonont rotifers. J. Hered. 104, 357–370.
Harring, H.K., Myers, F.J., 1922. The rotifer fauna of Wisconsin. Trans. Wisc.
Academy Sci., Arts Lett. 20, 553–662.
Hochberg, R., 2006. On the serotonergic nervous system of two planktonic
rotifers, Conochilus coenobasis and C. dossuarius (Monogononta,
Flosculariacea, Conochilidae). Zool. Anz. 245, 53–62.
270
Hochberg, R., 2009. Three-dimensional reconstruction and neural map of
the serotonergic brain of Asplanchna brightwellii (Rotifera, Monogononta). J. Morphol. 270, 430–441.
Hochberg, R., Ablak Gurbuz, O., 2008. Comparative morphology of
the somatic musculature in species of Hexarthra and Polyarthra
(Rotifera, Monogononta): its function in appendage movement and
escape behavior. Zool. Anz. 247, 233–248.
Hofmann, W., 1982. On the coexistence of two pelagic Filinia species
(rotatoria) in lake plußee i. Dynamics of abundance and dispersion.
Arch. für Hydrobiol. 95, 125–137.
Hovius, J.T., Beisner, B.E., McCann, K.S., Yan, N.D., 2007. Indirect food
web effects of Bythotrephes invasion: responses by the rotifer Conochilus in Harp Lake, Canada. Biol. Invasions 9, 233–243.
Hudson, C.T., Gosse, P.H., 1886. The Rotifera; or Wheel-animalcules,
Both British and Foreign, vols I and II, Longmans, Green, and Co.,
London.
Hyman, L.H., 1951. The Invertebrates: Acanthocephala, Aschelminthes,
and Entoprocta. The Pseudocoelomate Bilateria, vol. III, McGrawHill, New York, NY. 572 pp.
Ignoffo, T.R., Bollens, S.M., Bochdansky, A.B., 2005. The effect of thin
layers on the vertical distribution of the rotifer Brachionus plicatilis. J.
Exp. Mar. Biol. Ecol. 316, 167–181.
Iltis, A., Riou-Duvat, S., 1971. Variations saisonniéres du peuplement en
rotifères des eaux natronées du Kanem (Tchad). Cah. Orstom Hydrobiol. 5, 101–112.
Jersabek, C.D., Bolortsetseg, E., Taylor, H.L., 2010. Mongolian rotifers on
microscope slides: instructions to permanent specimen mounts from
expedition material. Mong. J. Biol. Sci. 8, 51–57.
Jersabek, C.D., Segers, H., Dingmann, B.J., 2003. The Frank J. Myers
Rotifer collection at the Academy of Natural Sciences: the whole
collection in digital images. Acad. Nat. Sci. Phila. Special
Publication 20.
Jones, B.L., VanLoozen, J., Kim, M.H., Miles, S.J., Dunham, C.M.,
Williams, L.D., Snell, T.W., 2013. Stress granules from in Brachionus
majavacas (Rotifera) in response to a variety of stressors. Comp. Biochem. Physiology, Part A 116, 375–384.
King, C.E., 1983. A re-examination of the Lansing effect. Hydrobiologia
104, 135–139.
Kleinow, W., Röhrig, A., 1995. Enzyme activity measurement on isolated
organs of Brachionus plicatilis (Rotifera). Hydrobiologia 313/314,
171–174.
Koste, W., Shiel, R.J., 1987. Rotifera from Australian inland waters. II.
Epiphanidae and Brachionidae (Rotifera: Monogononta). Invertebr.
Taxon. 7, 949–1021.
Langley, J.M., Shiel, R.J., Nielsen, D.L., Green, J.D., 2001. Hatching from
the sediment egg-bank, or aerial dispersing? – the use of mesocosms in
assessing rotifer biodiversity. Hydrobiologia 446/447, 203–211.
Lawrence, C., Sanders, E., Henry, E., 2012. Methods for culturing saltwater rotifers (Brachionus plicatilis) for rearing larval zebrafish. Zebrafish 9, 140–146.
Lowe, C.D., Kemp, S.J., Bates, A.D., Montagnes, D.J.S., 2005. Evidence
that the rotifer Brachionus plicatilis is not an osmoconformer. Mar.
Biol. 146, 923–929.
Mark Welch, D.B., Ricci, C., Meselson, M., 2009. (Chapter 13) Bdelloid
Rotifers: progress in understanding the success of an evolutionary
scandal. In: Schön, I., Martens, K., van Dijk, P. (Eds.), Lost sex: The
evolutionary biology of parthenogenesis. Springer Science + Business
Media B.V, Dordrecht, pp. 259–279.
SECTION | III
Protozoa to Tardigrada
Melone, G., 2001. Rhinoglena frontalis (Rotifera, Monogononta): a scanning electron microscopic study. Hydrobiologia 466/467, 291–296.
Melone, G., Ricci, C., 1995. Rotatory apparatus in Bdelloids. Hydrobiologia 313/314, 91–98.
Molinero, J.C., Anneville, O., Souissi, S., Balvay, G., Gerdeaux, D., 2005.
Anthropogenic and climate forcing on the long-term changes of planktonic rotifers in Lake Geneva, Europe. J. Plankton Res. 28, 287–296.
Obertegger, U., Flaim, G., Sommaruga, R., 2008. Multifactorial nature of
rotifer water layer preferences in an oligotrophic lake. J. Plankton Res.
30, 633–643.
Obertegger, U., Smith, H.A., Flaim, G., Wallace, R.L., 2011. Using the
guild ratio to characterize pelagic rotifer communities. Hydrobiologia
662, 157–162.
Park, H.G., Lee, K.W., Cho, S.H., Kim, H.S., Jung, M.-M., Kim, H.-S.,
2001. High density culture of the freshwater rotifer, Brachionus
calyciflorus. Hydrobiologia 446/447, 369–374.
Pourriot, R., 1979. Rotiféres du sol. Rev. d’Ecologie Biol. Sol. 16, 279–312.
Pourriot, R., Francez, A.-J., 1986. Rotifères. Bull. Mens. la Société linnéenne Lyon 55, 2–37.
Pryakhin, A., Tryapitsina, G.A., Deryabina, L.V., Atamanyuk, N.I.,
Stukalov, P.M., Ivanov, I.A., Kostyuchenko, V.A., Akleyev, A.V.,
­
2012. Status of ecosystems in radioactive waste reservoirs of the
Mayak production association in 2009. Health Physcian 101, 61–63.
Ramulu, H.G., Raoult, D., Pontarotti, P., 2012. The rhizome of life: what
about metazoa? Front. Cell. Infect. Microbiol. 2, 50.
Ricci, C., Caprioli, M., 2005. Anhydrobiosis in bdelloid species, populations and individuals. Intergrative Comp. Biol. 45, 759–763.
Ricci, C., Caprioli, M., Fontaneto, D., Melone, G., 2007. Volume and morphology changes of a bdelloid rotifer species (Macrotrachela quadricornifera) during anhydrobiosis. J. Morphol. 269, 233–239.
Ricci, C., Fontaneto, D., 2003. Mediterranean rotifers: a very inconspicuous taxon. Biogeographia 25, 161–167.
Ricci, C., Melone, G., 1998. Dwarf males in monogonont rotifers. Aquat.
Ecol. 32, 361–365.
Ricci, C., Perletti, F., 2006. Starve and survive: stress tolerance and lifehistory traits of a bdelloid rotifer. Funct. Ecol. 20, 340–346.
Ruttner-Kolisko, A., 1974. Planktonic rotifers: biology and taxonomy. Die
Binnengewässer Suppl. 26, 1–146.
Santos-Medrano, G.E., Rico-Martinez, R., Velásquez-Rojas, C.A., 2001.
Swimming speed and Reynolds numbers of eleven freshwater rotifer
species. Hydrobiologia 446/447, 35–38.
Sarma, S.S.S., Nandini, S., 2007. Small prey size offers immunity to predation: a case study on two species of Asplanchna and three brachionid prey (Rotifera). Hydrobiologia 593, 67–76.
Schöll, K., Szövényi, G., 2011. Planktonic rotifer assemblages of the Danube River at Budapest after the red sludge pollution in Hungary. Bull.
Environ. Contam. Toxicol. 87, 124–128.
Schröder, T., 2003. Precopulatory mate guarding and mating behaviour
in the rotifer Epiphanes senta (Monogononta: Rotifera). Proc. Royal
Soc. Lond. B 270, 1965–1970.
Schröder, T., 2005. Diapause in monogonont rotifers. Hydrobiologia 546,
291–306.
Schröder, T., Howard, S., Arroyo, L., Walsh, E.J., 2007. Sexual reproduction and diapause of Hexarthra sp. (Rotifera) in short-lived Chihuahuan Desert ponds. Freshwater Biol. 52, 1033–1042.
Seaman, M.T., Gophen, M., Cavari, B.Z., Azoulay, B., 1986. Brachionus
calyciflorus Pallas as agent for removal of E. coli in sewage ponds.
Hydrobiologia 135, 55–60.
Chapter | 13
Phylum Rotifera
Segers, H., 2002. The nomenclature of the Rotifera: annotated checklist of
valid family and genus-group names. J. Nat. Hist. 36, 631–640.
Segers, H., 2007. Annotated checklist of the rotifers (Phylum Rotifera),
with notes on nomenclature, taxonomy and distribution. Zootaxa
1564, 1–104.
Segers, H., 2008. Global diversity of rotifers (Rotifera) in freshwater.
Hydrobiologia 595, 49–59.
Sendacz, S., Caleffi, S., Santos-Soares, J., 2006. Zooplankton biomass
of reservoirs in different trophic conditions in the state of São Paulo,
Brazil. Braz. J. Biol. 66, 337–350.
Serra, M., Snell, T.W., 2009. (Chapter 14) Sex loss in monogonont rotifers.
In: Schön, I., Martens, K., van Dijk, P. (Eds.), Lost sex: The evolutionary biology of parthenogenesis. Springer Science + Business Media,
B.V, Dordrecht, pp. 281–294.
Serra, M., Snell, T., Gilbert, J.J., 2005. Delayed mixis in rotifers: an adaptive response to the effects of density dependent sex on population
growth. J. Plankton Res. 27, 37–45.
Smith, H.A., Burns, A.R., Shearer, T.L., Snell, T.W., 2012. Three heat
shock proteins are essential for rotifer thermotolerance. J. Exp. Mar.
Biol. Ecol. 413, 1–6.
Smith, J.M., Cridge, A.G., Dearden, P.K., 2010. Germ cell specification
and ovary structure in the rotifer Brachionus plicatilis. EvoDevo
1, 5.
Snell, T.W., 1986. Effects of temperature, salinity and food level on sexual
and asexual reproduction in Brachionus plicatilis (Rotifera). Mar.
Biol. 92, 157–162.
Snell, T.W., 1998. Estimating chronic toxicity using rotifers. Stand. Methods Exam. Water Wastewater 20 (8420), 8–62. 68–65.
Snell, T.W., 2011. A review of the molecular mechanisms of monogonont
rotifer reproduction. Hydrobiologia 662, 89–97.
Snell, T.W., Garman, B.L., 1986. Encounter probabilities between male
and female rotifers. J. Exp. Mar. Biol. Ecol. 97, 221–230.
Sørensen, M.V., Giribet, G., 2006. A modern approach to rotiferan phylogeny: combining morphological and molecular data. Mol. Phylogenet.
Evol. 40, 585–608.
Starkweather, P.L., 1987. Rotifera. In: Pandian, T.J., Vernberg, F.J. (Eds.),
Animal energetics. Protozoa through insecta, Vol. 1. Academic Press,
Orlando, FL, pp. 159–183.
Stemberger, R.S., Gilbert, J.J., 1985. Body size, food concentration, and
population growth in planktonic rotifers. Ecology 66, 1151–1159.
Strojsová, M., Vrba, J., 2007. Rotifer digestive enzymes: direct detection
using the ELF technique. Hydrobiologia 593, 159–165.
Thorp, J.H., Mantovani, S., 2005. Zooplankton of turbid and hydrologically dynamic prairie rivers. Freshwater Biol. 50, 1474–1491.
271
Verschoor, A.M., Van der Stap, I., Helmsing, N.R., Lu¨rling, M., Van
Donk, E., 2004. Inducible colony formation within the Scenedesmaceae: adaptive responses to infochemicals from two different herbivore taxa. J. Phycol. 40, 808–814.
Wallace, R.L., 1980. Ecology of sessile rotifers. Hydrobiologia 73,
181–193.
Wallace, R.L., Edmondson, W.T., 1986. Mechanism and adaptive significance of substrate selection by a sessile rotifer. Ecology 67, 314–323.
Wallace, R.L., 1987. Coloniality in the phylum Rotifera. Hydrobiologia
147, 141–155.
Wallace, R.L., Ricci, C., 2002. (Chapter 2) Rotifera. In: Rundle, S.D.,
­Robertson, A.L., Schmid-Araya, J.M. (Eds.), Freshwater Meiofauna:
Biology and ecology. Backhuys Publishers, Leiden, pp. 15–44.
Wallace, R.L., Smith, H.A., 2009. Rotifera. In: Likens, G.E. (Ed.), Encyclopedia of inland waters. Elsevier, Oxford, pp. 689–703.
Wallace, R.L., Snell, T.W., Ricci, C., Nogrady, T., 2006. Rotifera, second
ed. Biology, ecology and systematics, vol. 1, Backhuys Publishers,
Leiden. 299 pp.
Walsh, E.J., Salazar, M., Remirez, J., Moldes, O., Wallace, R.L., 2006. Predation by invertebrate predators on the colonial rotifer Sinantherina
socialis. Invertebr. Biol. 125, 325–335.
Walsh, E.J., Schröder, T., Wallace, R.L., Ríos Arana, J.V., Rico-Martínez, R.,
2008. Rotifers from selected inland saline waters in the Chihuahuan
Desert of México. Saline Syst. 4, 7.
Walsh, E.J., Zhang, L., 1992. Polyploidy and body size variation in a
natural population of the rotifer Euchlanis dilatata. J. Evol. Biol. 5,
345–353.
Weithoff, G., 2004. Vertical niche separation of two consumers (Rotatoria)
in an extreme habitat. Oecologia 139, 594–603.
Weithoff, G., 2007. Dietary restriction in two rotifer species: the effect of
the length of food deprivation on life span and reproduction. Oecologia 153, 303–308.
Weithoff, G., Wacker, A., 2007. The mode of nutrition of mixotrophic flagellates determines the food quality for their consumers. Funct. Ecol.
21, 1092–1098.
Wilson, C.G., Sherman, P.W., 2010. Anciently asexual bdelloid rotifers
escape lethal fungal parasites by drying up and blowing away. Science
327, 574–576.
Wilson, C.G., Sherman, P.W., 2013. Spatial and temporal escape from fungal parasitism in natural communities of anciently asexual bdelloid
rotifers. Proc. Royal Soc. B 280, 20131255.
Yoshida, T., Hairston Jr, N.G., Ellner, S.P., 2004. Evolutionary trade-off
between defence against grazing and competitive ability in a simple
­unicellular alga, Chlorella vulgarls. Proc. Royal Soc. B 271, 1947–1953.