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79
K.A. Pitt and C.H. Lucas (eds.), Jellyfi sh Blooms, DOI 10.1007/978-94-007-7015-7_4,
© Springer Science+Business Media Dordrecht 2014
Abstract Research on jellyfi sh blooms has focused heavily on the factors infl uencing
the production of blooms. Identifying the factors that cause blooms to collapse,
however, is important for predicting the duration of blooms and when they are likely
to disappear. We assembled studies from the literature to assess the persistence of
populations of medusae, the timing of the disappearance of the populations and the
potential cause of the populations’ declines. We found 76 observations that met our
criteria for inclusion that were derived from 33 studies and included 47 different
taxa. Most populations exhibited strongly seasonal patterns of occurrence, but the
population dynamics of the same or closely related species varied greatly across
small spatial and temporal scales. Duration of occurrence was negatively related to
latitude, but latitude explained only 8 % of the total variability, and no relationship
existed when tropical species were excluded from the analysis. Senescence after
spawning, infestations of parasites, food limitation, disease, low salinity, extreme water
temperatures, predation and intertidal stranding were most commonly cited as causing
blooms to collapse. Improving understanding of when and why blooms collapse will
benefi t coastal industries that are affected by blooms and greatly improve our under-
standing of how jellyfi sh blooms impact the ecology of the systems they inhabit.
Keywords Jellyfi sh blooms • Cnidarians • Biogeochemical cycling • Jelly-falls
• Population dynamics • Feeding ecology • Parasitism • Hyperiid amphipods
• Digenean trematodes • Physiological tolerance • Catabolism
Chapter 4
Bloom and Bust: Why Do Blooms
of Jellyfi sh Collapse?
Kylie A. Pitt , Ariella Chelsky Budarf , Joanna G. Browne ,
and Robert H. Condon
K. A. Pitt (*) • A. C. Budarf • J. G. Browne
Australian Rivers Institute and Griffi th School of Environment , Griffi th University ,
Gold Coast Campus , QLD 4222 , Australia
e-mail: k.pitt@griffi th.edu.au; a.budarf@griffi th.edu.au; jbrowne@museum.vic.gov.au
R. H. Condon
Dauphin Island Sea Lab , 101 Bienville Blvd , Dauphin Island , AL 36528 , USA
Department of Marine Science , University of South Alabama , Mobile , AL 36688 , USA
e-mail: rcondon@disl.org
80
4.1 Introduction
Jellyfi sh (i.e. cnidarian medusae and ctenophores) are renowned for their ‘boom and
bust’ population dynamics. Prolifi c rates of production, coupled with growth
rates (based on wet weight) that are two to three times those of non-gelatinous
pelagic taxa (Pitt et al. 2013 ), can result in the seemingly sudden appearance of
conspicuous, and often spectacular, population blooms. The biomass of blooms
regularly exceeds 10 t wet weight 100 m
−3 (Lilley et al. 2011 ). Typically, however,
blooms are short-lived, sustained for periods of weeks to months, after which the
populations disappear, often abruptly (i.e. collapse).
Concern regarding the perceived global increase in jellyfi sh blooms, coupled
with their potential negative ecological and socioeconomic impacts, has, over the
past two decades, seen a surge in studies that have tried to identify the causes of
blooms (see Condon et al. 2012 ). Identifying natural and anthropogenic causes of
blooms is important for predicting bloom events, developing potential management
or eradication strategies and forecasting how jellyfi sh populations may respond to
changing ocean conditions. However, such enormous fl uctuations in biomass
(both appearance and disappearance) are likely to have major infl uences on the
ecology of marine systems. For example, the disappearance of what is often the
dominant predator of zooplankton releases predation pressure on zooplankton
and may initiate trophic cascades. Jellyfi sh also provide shelter to juvenile fi sh
and invertebrates and, therefore, may infl uence recruitment and population dynamics
of such taxa (see Doyle et al., Chap. 5 ). The sudden disappearance of jellyfi sh also
has major implications for biogeochemical cycling because jellyfi sh turn over large
quantities of assimilated material as carbon-rich dissolved organic material, which
is shunted toward rapid uptake and respiration by specifi c microbial phylotypes
(Condon et al. 2011 ). Because microbial respiration converts potential food web
energy into a form that can only be utilised by autotrophs (i.e. carbon dioxide), this
detour of carbon represents a diversion of carbon away from higher trophic levels.
Following the collapse of blooms, therefore, the transfer of carbon to higher
trophic levels may be restored. Moreover, microbial respiration associated with
decomposition of medusae can create an oxygen demand that exceeds the rate of
oxygen resupply, resulting in localised hypoxia or anoxia (Pitt et al. 2009a ).
Consequently, understanding the causes of declines, the timing and locations where
blooms collapse and the ecological and biogeochemical consequences of bloom
collapses is equally as important as understanding the production of blooms.
Senescing jellyfi sh typically exhibit increased rates of physical damage, loads of
parasites and rates of infection (Mills 1993 ). The pattern of mortality varies little
among taxa and usually involves degeneration of the tentacles, oral structures and
gonads, reduced swimming ability and, fi nally, necrosis of the epithelial tissues of the
bell (Brewer 1989 ; Kikinger 1992 ). The fate of moribund jellyfi sh is poorly known,
but their specifi c density exceeds that of seawater and also living jellyfi sh (Yamamoto
et al. 2008 ) suggesting that they are likely to sink rapidly. The observation of largely
K.A. Pitt et al.
81
intact moribund jellyfi sh on the seafl oor, so-called jelly-falls (Lebrato et al. 2012 ),
supports this argument. Rapid sinking of medusae may accelerate regional rates of
carbon export from surface waters in open ocean regions (Yamamoto et al. 2008 ;
Lebrato and Jones 2009 ) and increase transfer effi ciency of the biological pump to
the deep sea (Billett et al. 2006 ). For example, Billett et al. ( 2006 ) observed in the
Arabian Sea the massive ‘jelly-fl ux’ of Crambionella orsini carcasses at 3,000 m
following a surface bloom, which contained an order of magnitude more carbon
than the total annual carbon fl ux as measured by sediment traps. Given the dearth of
long-term time series of jellyfi sh communities and their biogeochemical infl uences
in the open ocean (Condon et al. 2012 ), it is unclear on what spatial and temporal
scales jelly-falls occur and how they are linked to ‘boom and bust’ dynamics of
jellyfi sh blooms, although recent information suggests that jelly- falls are prevalent
in coastal areas and oligotrophic gyres (Yamamoto et al. 2008 ; Lebrato et al. 2012 )
and have occurred over geological timescales (Hagadorn et al. 2002 ; Condon et al.
2012 ). Similarly, information on how jelly-falls relate to carbon export processes
are sparse (but see Lebrato et al. 2013 ) but jellyfi sh size, density and shape, physical
advection and microbial decomposition (Riemann et al. 2006 ; Tinta et al. 2012 ;
Lebrato et al. 2013 ) are likely the primary driving factors infl uencing sinking of
jellyfi sh blooms (Lebrato et al. 2011 ).
The major objective of this chapter is to analyse the literature to elucidate
temporal and spatial trends in the persistence of jellyfi sh populations and to identify
the major causes of declines in jellyfi sh populations. The major drivers identifi ed as
causing blooms to collapse are then reviewed.
4.2 Literature Analysis
We searched the literature for studies of population dynamics to assess the persistence
of the population, timing of the disappearance of the population and the potential
cause of the population’s decline. Although ctenophores are usually considered
‘jellyfi sh’, they were not included in the analysis because their populations are
typically restocked annually from overwintering populations (Costello et al. 2006 )
indicating that at least some of the population is perennial. Only studies that
sampled medusae at intervals of less than 2 months and that sampled for ≥ 1 year,
or that sampled from before the initial appearance of the population and until after
the population had disappeared, were included. A linear regression was used to
test whether the duration of the occurrence of populations was related to latitude.
The duration of the population was determined by the period when medusae were
abundant (i.e. rare occurrences were excluded; Table 4.1 ).
Our analysis found 76 observations that met our criteria and included 44 obser-
vations of hydrozoans (including four siphonophores) and 32 observations of
scyphozoans (Table 4.1 ). The observations were derived from 33 studies and included
47 different taxa. Only eight observations were derived from the southern hemisphere
4 Bloom and Bust: Why Do Blooms of Jellyfi sh Collapse?
82
Table 4.1 Patterns of occurrence of medusae (and ephyrae when reported) and potential source of mortality for the population or for individuals within the
population. Lines indicate duration of occurrence. Dashed lines indicate that medusae or ephyrae were rare (as defi ned or stated by author). Blue lines = northern
hemisphere, green lines = southern hemisphere (seasons in the table refer to the appropriate season in the respective hemisphere). A single line per entry
indicates that only 1 year was sampled; patterns were identical between years or indicate a general pattern if ≥ 2 years sampled. Two lines for a given entry
represent differences in patterns of occurrence during each year
K.A. Pitt et al.
83
(continued)
4 Bloom and Bust: Why Do Blooms of Jellyfi sh Collapse?
84
Table 4.1 (continued)
Taxon: H hydrozoa, SP siphonophore, S scyphozoa
a Data were presented for two locations: Korsfjord and the nearby Fanafjord. To avoid over- representation of this one study in the table, only data for Korsfjord
are presented because the Korsfjord was sampled more extensively than the Fanafjord. Medusae were considered to be rare when densities were <10 ind. 1,000 m
−3
b Reported as Phialidium hemisphaerica
c Reported as Phialucium carolinae
d Invasive species
e Reported as Podocoryne minima
f Sampled Kiel Bight and Eckernforde Bay from May to Sept. Only data for Eckernforde Bay are included because the occurrence of the population in Kiel
Bight exceeded the duration of the sampling programme
g Mostly represented by ephyrae
K.A. Pitt et al.
85
(seven from Australia and one from Brazil). Two studies from Norway (Hosia and
Båmstedt 2007 ) and Portugal (Primo et al. 2012 ) which sampled multiple hydro-
zoan taxa contributed 17 and 9 observations, respectively. Most populations exhib-
ited strongly seasonal patterns with the majority exhibiting greatest abundances
between mid-spring and mid-autumn in their respective hemispheres. Seventeen
taxa occurred year-round, but at least one third of those still exhibited strong
seasonal variations in abundances. Rarely was the cause of the decline in the
population reliably identifi ed; however, authors frequently speculated about the
cause of mortality, which included senescence after spawning, infestations of parasites,
food limitation, disease, low salinity, extreme water temperatures (low and high),
predation, advection and intertidal stranding.
Surprisingly, populations of the same or closely related species sometimes
exhibited different dynamics at different locations or times. For example, the
most commonly sampled species, the scyphozoan Aurelia aurita , exhibited
strong seasonal patterns of occurrence in six studies but occurred throughout the
year in seven studies (Table 4.1 ). The pattern of occurrence of A. aurita also
varied substantially among years at a single location. For example, in Tomales Bay,
California, A. aurita exhibited a seasonal occurrence during 1 year, whilst the
population persisted throughout the following year (Hamner and Jenssen 1974 ).
Patterns of occurrence can also vary greatly over spatial scales of 10s of kilometres.
In the southern UK, for example, A. aurita persists year-round in a man-made
coastal lake but occurs seasonally nearby in Southampton Water (Lucas et al. 1997 ).
However, some caution must be applied to these observations of Aurelia because
the genus contains numerous cryptic species (Dawson and Jacobs 2001 ) and
some variations (particularly among locations) could refl ect taxonomic differences.
The invasive rhizostome, Phyllorhiza punctata , occurs predominantly during summer
and autumn in subtropical and temperature locations such as southern Western
Australia (Rippingale and Kelly 1995 ), the Gulf of Mexico (Graham et al. 2003 )
and southern Brazil (Haddad and Nogueira 2006 ), but populations of medusae
persist year-round in tropical Puerto Rico, despite still exhibiting distinct seasonal
cycles of recruitment and mortality (García 1990 ). These observations suggest that
populations of medusae may rarely achieve their potential maximum longevity
and that environmental conditions are most likely the primary drivers of mortality.
This conclusion is further supported by observations that medusae can survive
much longer (sometimes several years) in captivity than they do in the fi eld
(Zahn 1981 ).
The persistence of populations was negatively correlated with latitude ( P = 0.02),
but latitude explained only a small amount of the total variability (r
2 = 0.081), and
the relationship was largely driven by the year-round persistence of three of the
four tropical species recorded (i.e. Aurelia aurita and Mastigias papua in Palau and
Phyllorhiza punctata in Puerto Rico) (Fig. 4.1 ). When tropical species were
excluded, no relationship with latitude existed (r
2 = 0.006; P > 0.05).
4 Bloom and Bust: Why Do Blooms of Jellyfi sh Collapse?
86
4.3 Common Causes of Mortality of Medusae
4.3.1 Food Limitation
Medusae can be voracious predators of zooplankton. The very high water content of
medusae enables them to attain body sizes that are much larger than other plankti-
vores of equivalent carbon content (Acuña et al. 2011 ). This trait enables medusae
to support large feeding structures that can effi ciently clear large volumes of water
and theoretically enables medusae to survive in lower concentrations of prey than
other competing planktivores, such as fi sh (Acuña et al. 2011 ). Changes in growth
(Olesen et al. 1994 ) and biomass (Möller 1980 ; Miglietta et al. 2008 ) of medusae
populations often correlate with zooplankton production after correcting for tempo-
ral lags. When prey are plentiful, medusae can grow rapidly (e.g. wet weight-
specifi c growth of 0.88 d
−1 for Cotylorhiza tuberculata ; Kikinger 1992 ), and the
biomass of the population has the potential to accumulate until rates of predation by
medusae exceed secondary production of zooplankton and the biomass of medusae
cannot be sustained (Purcell and Decker 2005 ). When prey become limited, how-
ever, growth of medusae may be inhibited, and individuals may attain smaller sizes
than when food is unlimited (e.g. Schneider and Behrends 1994 ; Lucas et al. 1997 ).
Indeed, food limitation is regularly cited as a major cause of population declines
(Table 4.1 ). However, sometimes medusae have continued to grow (Møller and
Riisgård 2007 ) and accumulate biomass (Olesen et al. 1994 ) despite the biomass of
0
2
4
6
8
10
12
0 102030405060
Duration of occurrence (months)
Latitude (degrees N or S)
Tropics
Fig. 4.1 Relationship between duration of occurrence of medusae and latitude
K.A. Pitt et al.
87
zooplankton appearing to be too low to support the population. This may be because
rates of secondary production are very high despite the low standing biomass or that
medusae effi ciently exploit patches of zooplankton or may be able to supplement
their diet by feeding on picoplankton (e.g. cyanobacteria) and microplankton
(e.g. ciliates). Alternatively, the biomass of zooplankton in these studies may have
been underestimated because neither study sampled the demersal zooplankton that
emerges from the benthos into the water column at night. Emergent zooplankton are
an important dietary source for jellyfi sh because tactile predators such as jellyfi sh
can feed continuously during the day and night (Pitt et al. 2008 ), which may also
allow them to outcompete visual predators for food resources in waters that are
dark or contain high humic content (Aksnes et al. 2004 ). Indeed, in Kertinge Nor,
Denmark, densities of epibenthic copepods in the water column can be 20 times
greater at night than during the day (Olesen et al. 1994 ) and could, therefore, provide
a signifi cant food source.
If food is limited when medusae fi rst recruit, growth appears to be inhibited, and
medusae attain only small sizes. In Horsea Lake, UK, zooplankton productivity is
much lower than in the nearby Southampton Water, and A. aurita in Horsea Lake
are, correspondingly, much smaller than those in Southampton Water (Lucas et al.
1997 ). Moreover, the bell diameter of A. aurita was negatively correlated with pop-
ulation density over 20 years of observations in Kertinge Nor, Denmark (Riisgård
et al. 2010 ) and over 9 years in Kiel Bight, Germany (Schneider and Behrends
1994 ), suggesting that competition for food may limit growth.
4.3.2 Predation
Until recently, jellyfi sh were considered a trophic dead end; however, recent studies
indicate jellyfi sh are consumed by a variety of marine predators, including turtles,
birds, fi sh and other gelatinous zooplankton (reviewed by Arai 2005 ). While a
diverse range of predators feed on jellyfi sh, predation by fi sh and other gelatinous
zooplankton has the largest potential to impact jellyfi sh populations (Arai 2005 ).
Top-down regulation of jellyfi sh populations is diffi cult to demonstrate and quan-
tify; however, several authors have speculated that intense, intra-guild predation by
other gelatinous predators can regulate some medusae (Table 4.1 ). For example, in
Nova Scotia, Canada, the hydromedusa Rathkea octopunctata comprised 34 % of
the diet of the scyphomedusa Aurelia aurita indicating that A. aurita may regulate
natural populations of R. octopunctata (Matsakis and Conover 1991 ). In Norway,
Cyanea capillata preys heavily on A. aurita (Fig. 4.2 ), and the decline in the
A. aurita population coincides with an increase in C. capillata (Båmstedt et al. 1994 ).
Overlapping temporal succession of several hydrozoans in Norway may similarly
indicate intra-guild predation (Hosia and Båmstedt 2007 ).
A wide range of fi sh consume jellyfi sh, including spiny dogfi sh, chum
salmon, ocean sunfi sh, Atlantic mackerel and Atlantic cod (Arai
1988 ; Ates 1988 ;
Link and Ford 2006 ). Although fi sh probably exert signifi cant predatory pressure,
4 Bloom and Bust: Why Do Blooms of Jellyfi sh Collapse?
88
the importance of jellyfi sh as a dietary component is unknown due to unquantifi ed
digestion rates (Purcell and Arai 2001 ; Cardona et al. 2012 ). Diffi culties with gut
content analysis may be circumvented by using stable and enriched isotopes
(Pitt et al. 2009b ) and molecular techniques. A recent study in the Mediterranean
Sea used
13 C and
15 N stable isotopes as a tool to estimate the relative contribution of
gelatinous zooplankton to the diets of several apex predators (Cardona et al. 2012 ).
Although this study provided evidence that loggerhead sea turtles, ocean sunfi sh
and various opportunistic feeders potentially consume large quantities of jellyfi sh,
further research is needed to quantify rates and determine whether these predators
can regulate populations of jellyfi sh.
4.3.3 Parasitism
Parasitism is likely to be an important factor in the decline of many jellyfi sh blooms
and in the regulation of medusae populations. Medusae are infected by many types
of parasites, including hyperiid amphipods (Laval
1980 ; Dittrich 1988 ); digenean
trematodes, or fl ukes (Martorelli and Cremonte 1998 ); cestodes (Vannucci-Mendes
1944 ); isopods (Barham and Pickwell 1969 ); nematodes (Svendsen 1990 );
barnacles (Pagès 2000 ); sea anemones (McDermott et al. 1982 ) and, potentially,
microbes (Doores and Cook 1976 ). Parasites that infect non-gelatinous hosts
can cause the host populations to crash (e.g. krill: Gómez-Gutiérrez et al. 2003 ;
Fig. 4.2 Cyanea capillata capturing Aurelia aurita in Kiel Fjord, Germany (Reproduced by
permission of Kylie Pitt)
K.A. Pitt et al.
89
fi sh: Heins et al. 2010 ), and there is strong circumstantial evidence to suggest that
hyperiid amphipods may contribute to declines in medusa populations (Mills 1993 ),
and ctenophore populations have also been adversely affected by platyhelminth
worms (Yip 1984 ) and parasitic anemones (Reitzel et al. 2007 ). Blooms of jellyfi sh
are likely to be particularly susceptible to parasitism because abundances of para-
sites are positively correlated to densities of hosts (Arneberg et al. 1998 ) and the
population size of hosts is a determinant of parasite infection (Bagge et al. 2004 ).
4.3.3.1 Hyperiid Amphipods
Hyperiid amphipods are a paraphyletic group of marine amphipods whose features
(e.g. large eyes, maxillipeds with no palps) are believed to have arisen through their
association with planktonic hosts (Lützen 2005 ). While some hyperiid amphipods
are primarily free-living, most appear to depend on gelatinous hosts for at least
some stage of their life cycle (Arai 2005 ). These hosts include medusae (Fig. 4.3a ),
siphonophores, planktonic molluscs and salps (Gasca and Haddock 2004 ). In many
hyperiid species, females brood eggs and then deposit juveniles onto the host.
The juveniles then feed on their host until they reach a more independent stage
(Laval 1980 ). Some hyperiid adults continue to feed on their host’s tissues (Towanda
and Thuesen 2006 ), while others become free-living. Parathemisto gaudichaudi is
generally regarded as free-living; however, juveniles have been found associated
with salps (Madin and Harbison 1977 ). Other hyperiids attach to the outside of their
host and feed on plankton, entrained (Condon and Norman 1999 ) or caught by
the host (Laval 1972 ).
Hyperiid amphipods can be prevalent in populations of medusae. At times,
100 % of the population may be infected (Towanda and Thuesen 2006 ), and indi-
vidual medusae may host hundreds of hyperiids (Dittrich 1988 ; Towanda and
Thuesen 2006 ). Medusae have a remarkable ability to regenerate damaged tissues
when food is abundant (Mills 1993 ), but if dense infestations of parasites occur
during times when food is scarce, mortality may occur. For example, prior to the
disappearance of the hydromedusae Aequorea victoria and Mitrocoma cellularia
from Puget Sound, USA, individuals exhibited high proportions of grazing damage
(>75 % and 67–100 %, respectively) which was attributed primarily to the hyperiid
amphipods Parathemisto pacifi ca and Hyperia medusarum . Low proportions of
hydromedusae had food in their guts (44 % and 66 %, respectively) and were seem-
ingly unable to regenerate lost tissue. The hyperiid Hyperia galba had a similar
effect on populations of the scyphomedusae Chrysaora hysoscella , Aurelia aurita ,
Rhizostoma pulmo , Cyanea capillata and C. lamarckii over two consecutive years
in waters around Helgoland in the North Sea (Dittrich 1988 ). By autumn almost all
medusae were parasitised, and the number of amphipods per medusa reached 486
on C. hysoscella . The increasing rates of infection coincided with the medusae
shrinking, as the hyperiids consumed the gonads and then the mesoglea. Regeneration
by medusae appeared unable to offset rates of tissue loss, and by the end of autumn,
all the medusae had disappeared (Dittrich
1988 ). While there have been many
4 Bloom and Bust: Why Do Blooms of Jellyfi sh Collapse?
90
studies on hyperiid amphipods and their hosts (see reviews of Harbison et al. 1977 ;
Madin and Harbison 1977 ; Laval 1980 ), only Mills ( 1993 ) and Dittrich ( 1988 )
attempted to determine their effect on medusae populations. Other species of medu-
sae for which hyperiids may have caused or contributed to the disappearance of
populations include Aurelia aurita (Möller 1980 ; Møller and Riisgård 2007 ) and
Cyanea capillata (Metz 1967 ).
4.3.3.2 Digenean Trematodes
Digenean trematodes, which are parasitic fl atworms (fl ukes), infect at least 62 species
of medusae (Browne unpubl.). Although there are approximately 18,000 species
Fig. 4.3 ( a – b ) Ectoparasites
in Port Phillip Bay, Australia;
( a ) hyperiid amphipods
Hyperia gaudichaudi
(indicated with arrows ) on
the oral arms of the
scyphozoan Catostylus
mosaicus (scale bar is 2 cm)
and ( b ) anemone Peachia
hilli (indicated by arrow )
attached to the scyphozoan
Pseudorhiza haeckeli (scale
bar is 0.5 cm) (Reproduced
by permission of Joanna
Browne)
K.A. Pitt et al.
91
of digeneans, only 13 infect medusae (Browne unpubl.). Digeneans have a complex
life cycle, mostly involving three hosts. The fi rst host is normally a mollusc, the
intermediate host is normally another invertebrate and the fi nal host is almost
always a vertebrate. Different life history stages of the parasite occur in each host,
and some are capable of reproduction (e.g. sporocysts in the mollusc host and
sexual adults in the vertebrate host). Digeneans that use jellyfi sh as an intermediate
host leave their mollusc host and penetrate the jellyfi sh and develop into metacer-
cariae which is a juvenile resting stage in an intermediate host. When the jellyfi sh
are eaten by suitable fi sh hosts, the metacercariae develop into sexual adults within
the fi sh. The metacercariae are likely to feed upon the jellyfi sh tissue, and highly
parasitised medusae can have an ‘ablandamiento total’ (=overall softening) of tissue
(Girola et al. 1992 ).
The proportion of medusae infected by digeneans in a population can be very
high (Fraser 1970 ) and is often higher than that observed in other planktonic inter-
mediate hosts (Marcogliese 1995 ). Rates of infection by digeneans in studies that
sampled >1,400 individuals of one medusa species ranged from 0.1 % to 97.6 %
(Diaz Briz et al. 2012 ) and depended on the species of digenean and host and
season. The only study to have examined the direct effect of digenean parasites on
a population of gelatinous zooplankton has focused on ctenophores. Yip ( 1984 )
sampled populations of the host ctenophore Pleurobrachia pileus monthly for 3½
years and observed a sharp decline in abundance of the ctenophore following
periods of heavy infection by parasites (predominately Opechona bacillaris and
didymozoid larvae). She proposed that effects on the host could include competition
for food, consumption of body tissue and increasing body weight of the host interfering
with normal movement.
4.3.3.3 Parasitic Anemones
Larval anemones of the genera Edwardsiella and Peachia (Fig. 4.3b ) parasitise jelly-
fi sh and feed on their intestinal fl uids, gonads and mouth tissues (Badham 1917 ;
Spaulding 1972 ; Mills 1993 ). As adults, the anemones are benthic and free-living
(McDermott et al. 1982 ; Reitzel et al. 2006 ). While many medusae are infected by
larval anemones (Lauckner 1980 ), the only ecological studies about their effects on
host populations have been done on ctenophores. In the laboratory, larval E. lineata
decreased the growth rates of their host ctenophore M. leidyi and indirectly decreased
fecundity through their infl uence on host size (Bumann and Puls 1996 ). These par-
asite-induced effects led the anemone to be proposed as a biological control on its
invasive host M. leidyi (Bumann and Puls 1996 ). However, using the anemone as a
biological control would be risky because the anemone is linked to the skin irritation
‘sea bathers eruption’ (Freudenthal and Joseph 1993 ) and may alter benthic
communities (Bumann and Puls 1996 ). Recently, E. lineata is believed to have
followed its invasive host to the northeast Atlantic (although there is some diffi culty
in differentiating E. lineata and the similar E. carnea ) (Selander et al.
2010 ).
4 Bloom and Bust: Why Do Blooms of Jellyfi sh Collapse?
92
4.3.3.4 Importance of Medusae Parasites and Relevance to Blooms
While there have been many studies of some medusae parasites, particularly
hyperiids, those above are examples of the few that have examined the effects
of parasites on medusae populations through time. Many medusa parasites
(e.g. microbes) that have the potential to have detrimental effects on their hosts are
poorly understood (Ohtsuka et al. 2009 ), but further research will enable perspective
of their importance and relevance to medusae mortality and decline of blooms.
Recently the ability of parasites to affect entire communities has been high-
lighted (Lafferty 2008 ; Hatcher et al. 2012 ). Medusae parasites may infl uence
other organisms through predation, transference and regulation of host populations.
Parasites which use medusae as intermediate hosts may be transferred to commer-
cially important species. For example, the mackerels Scomber scombrus , S. japonicus
and S. australasicus are infected by numerous digeneans that use jellyfi sh as hosts
(Bray and Gibson 1990 ; Bartoli and Bray 2004 ). The transmission of parasites
depends on the density of the hosts (e.g. farmed salmonids and sea lice: Jansen
et al. 2012 ), and when jellyfi sh form blooms, parasites such as larval anemones and
hyperiids may spread more easily between medusae hosts (Spaulding 1972 ; Laval
1980 ). There may also be increased transfer of parasites to predators; the parasitic
anemone, Edwardsiella lineata , is transferred when its host Mnemiopsis leidyi is
eaten by Beroë ovata (Reitzel et al. 2007 ). Peaks in medusa populations offer an
increased abundance of hosts and therefore appear to be linked to peaks in parasite
abundance (Williams and Robins 1981 ; Dittrich 1988 ). Medusae parasites may
have positive effects on other animals by relieving predation pressure by the medusae
hosts, or they may be a food source. For example, hyperiid amphipods are picked
directly from their hosts by pile perch Rhacochilus vacca , the symbiont crab Cancer
gracilis (Towanda and Thuesen 2006 ) and sea birds (Harrison 1984 ).
4.3.4 Disease
While disease is often considered to be a potential cause of mortality in medusae,
few studies have confi rmed infections as a cause of death. Hydromedusae with
bacterial infections are able to recover if suffi cient food is available (Mills 1993 ).
However, at the end of the hydromedusae’s seasonal occurrence, the reduced avail-
ability of prey may render them more susceptible to these infections (Mills 1993 ).
Late in the season, for example, over 80 % of the hydromedusa Clytia gregaria had
bacterial infections on their bells, which was thought to contribute to mortality
when coupled with limited food availability (Mills 1993 ). Similarly, mortality of
Gonionemus vertens was thought to be primarily due to infection characteristically
associated with senescence (Mills 1993 ). Bacteria also infected wounds generated
by bites of argonauts in the rhizostome Phyllorhiza punctata which may have
exacerbated the physical injuries incurred (Heeger et al.
1992 ). Although pathogens
other than bacteria (e.g. viruses and fungi) probably infect medusae, no studies of
such pathogens exist.
K.A. Pitt et al.
93
4.3.5 Death Post-Spawning
Scyphozoan jellyfi sh have, on several occasions, been observed to die shortly after
spawning (Table 4.1 ). Mortality post-spawning has been examined particularly in
Aurelia aurita . In the Baltic Sea mortality rates prior to maturation were low, but
after spawning the medusae degraded and died (Möller 1980 ). Starvation and
increased parasitism were suggested to be the major cause of degradation rather
than spawning itself. However, Spangenberg ( 1965 ) observed that sexual products
and gastric fi laments of Aurelia aurita were released simultaneously during spawn-
ing. Because gastric fi laments (or gastric cirri) are necessary for digestion within
the stomach, their loss during spawning suggests starvation as the most likely
explanation for deterioration in this case (Spangenberg 1965 ; Arai 1997 ). Contrary
to these studies, however, Hamner and Jenssen ( 1974 ) found that after spawning
medusae were able to ‘ripen’ gonads within a couple of weeks. Therefore, their
observations in the laboratory did not support simultaneous deterioration of somatic
and reproductive tissue, although they did observe deterioration and mortality
post-spawning in the fi eld.
Cyanea is another genus that reportedly spawns and then deteriorates (Fancett
1986 ). However, in the Niantic River estuary, USA, Cyanea sp. lose their tentacles
prior to losing their oral folds (which contain the planulae) and gonads, and so the
major cause of death may be starvation due to loss of tentacles rather than spawning
(Brewer 1989 ).
4.3.6 Metabolic Intolerances to Physical Conditions
Patterns of occurrence of many medusae are often correlated with seasonal changes
in physical parameters such as temperature or salinity (e.g. Fancett 1986 ; Lo and
Chen 2008 ; Primo et al. 2012 ) which, in turn, are correlated with a variety of other
changes, such as decreased zooplankton production. Only in regions where seasonal
changes in the physical environment exceed the physiological tolerances of species,
however, are physical factors likely to be the main driver of mortality. To rigorously
identify physical conditions as the main cause of mortality requires experiments on
tolerance limits to be undertaken, preferably at the location of interest to account for
local adaptation, and then related to fi eld observations.
4.3.6.1 Temperature
Although populations of medusae often disappear when water temperatures decrease
during autumn, only in Chesapeake Bay is there robust evidence that death of medusae
is caused by cooling water temperatures. In laboratory experiments, the pulsation
rate of Chrysaora quinquecirrha slows with declining water temperature, and at
10 °C medusae cease to pulse and die (Gatz et al. 1973 ). These results are consistent
4 Bloom and Bust: Why Do Blooms of Jellyfi sh Collapse?
94
with observations in the fi eld and laboratory, whereby medusae sink deeper into the
water column when the temperature decreases to 15 °C and then disappear entirely
from the water column at 10 °C (Sexton et al. 2010 ), suggesting that the cold water
may have been the dominant cause of death. Warm temperatures can also invoke
mortality. For example, following an ENSO event in 1997–1998 which elevated
temperatures 1–2 °C above their long-term seasonal average, the normally perennial
population of the zooxanthellate rhizostome Mastigias papua disappeared from
Ongeim’l Tketau lake in Palau (Dawson et al. 2001 ; Martin et al. 2006 ). Concurrent
laboratory experiments showed that mortality of medusae increased greatly at
temperatures similar to those measured in the lake; therefore, warm water was
considered the major cause of mass mortality (Dawson et al. 2001 ).
4.3.6.2 Salinity
Evidence linking changes in salinity to mortality events of medusae is relatively
weak and constrained to correlative observations. For example, Aurelia aurita
disappears from the surface waters of a coastal lagoon in Taiwan following heavy
rain during summer, but it is unclear whether the population dies, is advected from
the lagoon or simply remains below the halocline (Lo and Chen 2008 ). In Western
Australia, the distribution and persistence of Phyllorhiza punctata appears to be
correlated to rainfall, with periods of heavy rain preceding the disappearance of the
population (Rippingale and Kelly 1995 ). Populations of Chrysaora quinquecirrha
in the mesohaline region of Chesapeake Bay are similarly correlated with stream-
fl ow and salinity (Cargo and King 1990 ; Purcell et al. 1999 ), but research has
focused mainly on the effects of salinity on production of medusae rather than as a
cause of mortality.
4.3.6.3 UV Radiation
Ultraviolet (UV) radiation damages tissues and induces vertical migration in zoo-
plankton (Rhode et al. 2001 ). Consequently UV radiation could be detrimental to
medusae. In Lake Tanganyika, the freshwater hydrozoan, Limnocnida tanganjicae ,
died within 1 h when exposed to UV radiation equivalent to that found close to the
surface waters (Salonen et al. 2012 ). However, L. tanganjicae undertakes diel verti-
cal migration, and this, presumably, prevents mortality in situ. The upside-down
jellyfi sh Cassiopea sp. is restricted to occurring in shallow waters due to its need to
photosynthesise and, therefore, may be susceptible to exposure to UV radiation.
The zooxanthellae within this species synthesise mycosporine-like amino acids
that have a photoprotective function and that can be translocated to the host to
provide protection against UV radiation (Banaszak and Trench 1995 ). Pigments
may also be formed through uptake of glycoproteins, which may serve to protect
cells in zooxanthellate medusae from UV radiation (Blanquet and Phelan
1987 ).
K.A. Pitt et al.
95
Whilst no studies have attributed large-scale mortality of medusae to UV radiation,
increasing levels of radiation could, potentially, induce mortality in shallow systems
where vertical migration is not possible.
4.3.7 Stranding
Mass strandings of jellyfi sh are common on beaches (e.g. Houghton et al. 2007 ;
Fuentes et al. 2010 ; Fig. 4.4 ) and, because of their conspicuous nature, often attract
the attention of media (Lilley et al. 2009 ; Condon et al. 2012 ). Strandings, however,
are more likely to be a consequence, rather than a cause of mortality for medusae,
and the timing of events may relate to oceanographic and weather conditions. For
example, large numbers of the rhizostome Cotylorhiza tuberculata strand on
beaches in Vlyho Bay, Greece, during autumn, associated with strengthening winds
(Kikinger
1992 ). These strandings may be facilitated by reduced swimming ability
associated with sloughing of the subumbrella muscles as the medusae senesce
(Kikinger 1992 ). Moreover, Chrysaora hysoscella that wash ashore on beaches in
the Irish Sea often lack peripheral tentacles and oral arms, indicating that these
medusae may have senesced prior to stranding (Houghton et al. 2007 ). Mass strandings
of decaying medusae on beaches may represent a substantial input of carbon to
beach environments, which are typically poorly productive and rely on allochthonous
inputs of organic matter.
Fig. 4.4 Mass stranding of Crambione mastigophora at Cable Beach, Broome, Western Australia
(Reproduced by permission of James Browne, Kimberley Marine Research Station, Cygnet Bay)
4 Bloom and Bust: Why Do Blooms of Jellyfi sh Collapse?
96
4.4 Factors That Promote Survival of Jellyfi sh
The persistence of medusae populations may relate to their variable abilities to
either withstand the drivers of mortality or to recover from them (see also Lucas and
Dawson, Chap. 2 ). The ability of medusae to catabolise their own tissues when
starving and to heal wounds and regenerate lost body parts is likely to provide
medusae with the ability to potentially survive stressors.
4.4.1 Ability to Shrink When Starved
When food is scarce, most organisms can utilise stores of lipids to sustain
themselves. Medusae, however, contain approximately half the lipid content (as %
ash- free dry weight, AFDW) of non-gelatinous pelagic taxa (Clarke et al. 1992 ;
Donnelly et al. 1994 ), and the majority of lipids are phospholipids which constitute
components of cell membranes (Arai et al. 1989 , Costello 1992 ). Due to the lack of
storage lipids, the ubiquitous responses of medusae to starvation are to catabolise
their own tissues and rapidly lose mass (Hatai 1917 ; Hamner and Jenssen 1974 ;
Arai et al. 1989 ). The degree of degrowth can be remarkable. For example, Cassiopea
can lose up to 99 % of its mass (Mayer 1914 ), and A. aurita can shrink to a quarter
of its original diameter and remain viable; however, once the diameter is less than
2 cm, the medusae usually become deformed and deteriorate (Hamner and Jenssen
1974 ). In the hydromedusa Aequorea victoria , proteins, lipids and carbohydrates
are catabolised at similar rates (Arai et al. 1989 ). However, while A. aurita and
A. victoria shrink rapidly when starved (Hamner and Jensen 1974 ; Arai et al. 1989 ),
the hydromedusa Cladonema californicum actually increases diameter and maintains
an enlarged diameter for up to 28 days following the onset of starvation, despite
losing 69–77 % of its dry mass (Costello 1998 ). Maintaining their diameter
whilst losing mass, however, compromises their ability to swim (Costello 1998 ).
The difference in response of the few taxa for which starvation has been studied
may refl ect differences in their feeding ecologies. Specifi cally, C. californicum is an
ambush ‘sit and wait’ predator that relies on maximising encounter rates to capture
prey, whereas A. aurita and A. victoria are cruising predators that use vortices
generated by active swimming to entrain their prey (Costello 1998 ). Consequently,
A. aurita and A. victoria depend much more heavily on swimming to capture prey and
regrow. Maximising bell diameter, potentially at the expense of maintaining other
structures, such as muscles, may optimise survival of ambush predators and maximise
their chance for recovery once prey become more numerous (Costello 1998 ).
Aurelia aurita and Cladonema californicum can both regrow following more
than 6 weeks of starvation (Hamner and Jenssen 1974 ; Costello 1998 ). In both species
the pattern of growth following starvation is normal, and individuals can reinstate
normal feeding and reproductive processes. However, whilst in the laboratory
medusae exhibit an extraordinary ability to degrow and regrow, we could fi nd no
examples of cohorts of medusae recovering after shrinking in the fi eld. In the fi eld,
K.A. Pitt et al.
97
degrowth is usually determined from a decrease in the average size of medusae
(Möller 1980 ); however, decreases in average size can also be explained by selec-
tive mortality or advection of the larger size classes in the population (Brewer 1989 ;
Olesen et al. 1994 ) and, therefore, need to be interpreted cautiously. Degrowth
(where it has been claimed) is usually observed during autumn (e.g. Möller 1980 ;
Ishii and Båmstedt 1998 ; Møller and Riisgård 2007 ) which coincides with cooling
water temperatures and reduced rates of zooplankton production, conditions that
typically persist for several months. Whilst medusae can sustain at least three
months starvation in the laboratory (Hamner and Jensen 1974 ), the two studies of
regrowth by medusae have been undertaken at relatively warm and constant
temperatures (16–18 °C, Hamner and Jensen 1974 ; 18 °C, Costello 1998 – both
studies done in California). Indeed, interactive effects between regrowth and
temperature are yet to be tested but may demonstrate that regrowth is not viable
when water temperatures approach the thermal minimum for a species.
4.4.2 Ability to Heal Injuries and Regenerate Lost Body Parts
Medusae have remarkable abilities to heal injuries and regrow damaged body parts
(Zeleney 1907 ; Mills 1993 ). For example, parasitic hyperiid amphipods often consume
the manubria of the hydromedusa Aequorea victoria (Mills 1993 ). However, if the
damaged individual is transferred to an aquarium and fed well, it can regenerate a new
manubrium within 6 days (Mills 1993 ). Similarly, a hole penetrating the centre of the
umbrella of Mitrocoma can heal within 7 days (Mills 1993 ). Whilst injuries can heal
under laboratory conditions, recovery from injury also appears to occur in the fi eld.
For example, it is common to see substantial scars created by the healing of injuries
derived from the blades of boat propellers in large medusae (Pitt pers. obs.). Rates
of regeneration increase with severity of the injury, up until a threshold. For example,
regeneration of the oral arms of Cassiopea xamachana increased as additional oral
arms were removed, with the maximum rate of regeneration associated with the
removal of 6 of the 8 oral arms (Zeleney 1907 ). Moreover, jellyfi sh can also regen-
erate the same body parts multiple times (Zeleney 1907 ). Mechanisms of wound
healing are, however, very poorly studied. Very small wounds (1.2 mm diameter)
in the myoepithelial cells of the swimming muscle are closed by the muscle cells
differentiating into epithelial cells and migrating to the centre of the wound before
dedifferentiating into contractile muscle cells again (Lin et al. 2000 ).
4.5 Conclusions
Rarely have the causes of mortality of medusae been reliably identifi ed. Extreme
variability in persistence of populations of the same species among locations and
between years indicates that medusae may only rarely attain their maximum
4 Bloom and Bust: Why Do Blooms of Jellyfi sh Collapse?
98
physiological longevity in the fi eld, with environmental parameters that vary both
temporally and spatially the main drivers of mortality. Mortality is likely due to
multiple stressors interacting rather than individual events. Small variations in the
timing or magnitude of the stressors may invoke changes in the rate or timing of
mortality. Mass mortality, particularly in shallow or enclosed water bodies, such as
coastal lagoons and fjords, can have major implications for the ecology and bio-
geochemical cycling of the systems. Being able to predict the duration of blooms
and when they are likely to decline could benefi t coastal industries, such as tourism,
fi sheries and power generation, which are often negatively impacted by jellyfi sh
(see Lucas et al., Chap. 6 ). Reliable identifi cation of the factors leading to the
collapse of blooms should, therefore, be a priority for research.
References
Acuña JL, López-Urrutia Á, Colin S (2011) Faking giants: the evolution of high prey clearance
rates in jellyfi shes. Science 333:1327–1629
Aksnes DL, Nejstgaard J, Sœdberg ST (2004) Optical control of fi sh and zooplankton populations.
Limnol Oceanogr 49:233–238
Albert DJ (2005) Reproduction and longevity of Aurelia labiata in Roscoe Bay, a small bay on the
Pacifi c coast of Canada. J Mar Biol Assoc UK 85:575–581
Arai MN (1988) Interactions of fi sh and pelagic coelenterates. Can J Zool 66:1913–1927
Arai MN (1997) A functional biology of Scyphozoa. Chapman and Hall, London
Arai MN (2005) Predation on pelagic coelenterates: a review. J Mar Biol Assoc UK 85:523–536
Arai MN, Ford JA, Whyte JNC (1989) Biochemical composition of fed and starved Aequorea
victoria (Murbach et Shearer, 1902) (Hydromedusa). J Exp Mar Biol Ecol 127:289–299
Arneberg P, Skorping A, Grenfell B, Read AF (1998) Host densities as determinants of abundance
in parasite communities. Proc Roy Soc Lond Ser B 265:1283–1289
Ates R (1988) Medusivorous fi shes, a review. Zool Meded 62:29–42
Attrill MJ, Thomas RM (1996) Long-term distribution patterns of mobile estuarine invertebrates
(Ctenophora, Cnidaria, Crustacea: Decapoda) in relation to hydrological parameters. Mar Ecol
Prog Ser 143:25–36
Badham C (1917) On a larval actinian parasitic in a rhizostome. Q J Microsc Sci 62:221–229
Bagge A, Poulin R, Valtonen E (2004) Fish population size, and not density, as the determining
factor of parasite infection: a case study. Parasitology 128:305–313
Båmstedt U, Martinussen MB, Matsakis S (1994) Trophodynamics of the two scyphozoan jellyfi shes,
Aurelia aurita and Cyanea capillata , in western Norway. ICES J Mar Sci 51:369–382
Banaszak AT, Trench RK (1995) Effects of ultraviolet (UV) radiation on marine microalgal
invertebrate symbioses. II. The synthesis of mycosporine-like amino acids in response to
exposure to UV in Anthopleura elegantissima and Cassiopea xamachana. J Exp Mar Biol Ecol
194:233–250
Barham EG, Pickwell GV (1969) The giant isopod, Anuropus : a scyphozoan symbiont. Deep-Sea
Res 16:525–529
Bartoli P, Bray R (2004) Cephalolepidapedon saba ; Yamaguti, 1970 (Digenea: Lepocreadiidae),
the redescription of a species newly reported in European waters. Syst Parasitol 58:189–198
Barz K, Hirche H-J (2007) Abundance, distribution and prey composition of scyphomedusae in the
southern North Sea. Mar Biol 151:1021–1033
Batistić M, Jasprica N, Carić M, Lučić D (2007) Annual cycle of the gelatinous invertebrate
zooplankton of the eastern South Adriatic coast (NE Mediterranean). J Plankton Res
29:671–696
K.A. Pitt et al.
99
Billett DSM, Bett BJ, Jacobs CL, Rouse IP, Wigham BD (2006) Mass deposition of jellyfi sh in the
deep Arabian Sea. Limnol Oceanogr 51:2077–2083
Blanquet RS, Phelan MA (1987) An unusual blue mesogleal protein from the mangrove jellyfi sh
Cassiopea xamachana . Mar Biol 94:423–430
Bray R, Gibson D (1990) The Lepocreadiidae (Digenea) of fi shes of the north-east Atlantic: review
of the genera Opechona Looss, 1907 and Prodistomum Linton, 1910. Syst Parasitol
15:159–202
Brewer RH (1989) The annual pattern of feeding, growth, and sexual reproduction in Cyanea
(Cnidaria: Scyphozoa) in the Niantic River estuary, Connecticut. Biol Bull 176:272–281
Bumann D, Puls G (1996) Infestation with larvae of the sea anemone Edwardsia lineata affects
nutrition and growth of the ctenophore Mnemiopsis leidyi . Parasitology 113:123–128
Cardona L, de Quevedo IA, Borrell A, Aguilar A (2012) Massive consumption of gelatinous plank-
ton by Mediterranean apex predators. PLoS One 7:e31329
Cargo DG, King DR (1990) Forecasting the abundance of the Sea Nettle, Chrysaora quinquecir-
rha , in the Chesapeake Bay. Estuaries 13:486–491
Clarke A, Holmes LJ, Gore DJ (1992) Proximate and elemental composition of gelatinous
zooplankton from the Southern Ocean. J Exp Mar Biol Ecol 155:55–68
Condon RH, Norman MD (1999) Commensal associations between the hyperiid amphipod,
Themisto australis , and the scyphozoan jellyfi sh, Cyanea capillata . Mar Freshw Behav Physiol
32:261–267
Condon RH, Steinberg DK, del Giorgio PA, Bouvier TC, Bronk DA, Graham WM, Ducklow HW
(2011) Jellyfi sh blooms result in a major microbial respiratory sink of carbon in marine
systems. Proc Natl Acad Sci USA 108:10225–10230
Condon RH, Graham WM, Duarte CM, Pitt KA, Lucas CH, Haddock SHD, Sutherland KR,
Robinson KL, Dawson MN, Decker MB, Mills CE, Rhode JE, Malej A, Hermes M, Uye S-I,
Belcich S, Madin LP (2012) Questioning the rise of gelatinous zooplankton in the world’s
oceans. BioScience 62:160–169
Costello J (1991) Complete carbon and nitrogen budgets for the hydromedusa Cladonema califor-
nicum (Anthomedusa: Cladonemidae). Mar Biol 108:119–128
Costello JH (1998) Physiological response of the hydromedusa Cladonema californicum Hyman
(Anthomedusa: Cladonemidae) to starvation and renewed feeding. J Exp Mar Biol Ecol
225:13–28
Costello JH, Sullivan BK, Gifford DJ, Van Keuren D, Sullivan LJ (2006) Seasonal refugia,
shoreward thermal amplifi cation, and metapopulation dynamics of the ctenophore Mnemiopsis
leidyi in Narragansett Bay, Rhode Island. Limnol Oceanogr 51:1819–1831
Dawson MN, Jacobs DK (2001) Molecular evidence for cryptic species of Aurelia aurita (Cnidaria,
Scyphozoa). Biol Bull 200:92–96
Dawson MN, Martin LE, Penland LK (2001) Jellyfi sh swarms, tourists, and the Christ-child.
Hydrobiol 155:131–144
Decker MB, Brown CW, Hood RR, Purcell JE, Gross TFZ, Matanoski JC, Bannon RO,
Setzler- Hamilton EM (2007) Predicting the distribution of the scyphomedusa Chrysaora
quinquecirrha in Chesapeake Bay. Mar Ecol Prog Ser 329:99–113
Diaz Briz L, Martorelli S, Genzano G, Mianzan H (2012) Parasitism (Trematoda, Digenea) in
medusae from the southwestern Atlantic Ocean: medusa hosts, parasite prevalences, and
ecological implications. Hydrobiologia 690:215–226
Dittrich B (1988) Studies on the life cycle and reproduction of the parasitic amphipod Hyperia
galba in the North Sea. Helgol Mar Res 42:79–98
Donnelly J, Torres JJ, Hopkins TL, Lancraft TM (1994) Chemical composition of Antarctic
zooplankton during austral fall and winter. Polar Biol 14:171–183
Doores S, Cook TM (1976) Occurrence of Vibrio and other bacteria on the sea nettle, Chrysaora
quinquecirrha . Microb Ecol 3:31–40
Fancett M (1986) Species composition and abundance of scyphomedusae in Port Phillip Bay,
Victoria. Mar Freshw Res 37:379–384
4 Bloom and Bust: Why Do Blooms of Jellyfi
sh Collapse?
100
Fraser JH (1970) The ecology of the ctenophore Pleurobrachia pileus in Scottish waters. ICES J
Mar Sci 33:149–168
Freudenthal AR, Joseph PR (1993) Seabather’s eruption. New Engl J Med 329:542–544
Fuentes VL, Angel DL, Bayha KM, Atienza D, Edelist D, Bordehore C, Gilia J-M, Purcell JE
(2010) Blooms of the invasive ctenophore, Mnemiopsis leidyi , span the Mediterranean Sea in
2009. Hydrobiol 645:23–37
García JR (1990) Population dynamics and production of Phyllorhiza punctata (Cnidaria:
Scyphozoa) in Laguna Joyuda, Puerto Rico. Mar Ecol Prog Ser 64:243–251
Gasca R, Haddock SHD (2004) Associations between gelatinous zooplankton and hyperiid amphipods
(Crustacea : Peracarida) in the Gulf of California. Hydrobiologia 530–31:529–535
Gatz AJ, Kennedy VS, Mihursky JA (1973) Effects of temperature on activity and mortality of the
scyphozoan medusa, Chrysaora quinquecirrha . Chesapeake Sci 14:171–180
Girola CV, Martorelli SR, Sardella NH (1992) Presencia de metacercarias de Monascus fi liformis
(Digenea, Fellodistomidae) en hidromedusas del Océano Atlántico Sur (Presence of metacer-
cariae of Monascus fi liformis (Digenea, Fellodistomidae) in hydromedusae of the South
Atlantic Ocean). Rev Chil Hist Nat 65:409–415
Gómez-Gutiérrez J, Peterson WT, De Robertis A, Brodeur RD (2003) Mass mortality of krill
caused by parasitoid ciliates. Science 301:339
Graham WM, Martin DL, Felder DL, Asper VL, Perry HM (2003) Ecological and economic
implications of a tropical jellyfi sh invader in the Gulf of Mexico. Biol Invasions 5:53–69
Haddad MA, Hogueira M (2006) Reappearance and seasonality of Phyllorhiza punctata von
Lendenfeld (Cnidaria, Scyphozoa, Rhizostomeae) medusae in southern Brazil. Revista
Brasileira de Zoologia 23:824–831
Hagadorn JW, Dott RH Jr, Damrow D (2002) Stranded on a Late Cambrian shoreline: Medusae
from central Wisconsin. Geology 30:147–150
Hamner W, Jensen R (1974) Growth, degrowth, and irreversible cell differentiation in Aurelia
aurita . Am Zool 14:33–849
Hamner WM, Gillmer RW, Hamner PP (1982) The physical, chemical, and biological characteristics
of a stratifi ed, saline, sulfi de lake in Palau. Limnol Oceanogr 27:896–909
Harbison GR, Biggs DC, Madin LP (1977) Associations of Amphipoda Hyperiidea with gelatinous
zooplankton – II. Associations with Cnidaria, Ctenophora and Radiolaria. Deep-Sea Res
24:465–472
Harrison NM (1984) Predation on jellyfi sh and their associates by seabirds. Limnol Oceanogr
29:1335–1337
Hatai S (1917) On the composition of the medusa, Cassiopea xamachana and the changes in it
after starvation. Proc Natl Acad Sci USA 3:22–24
Hatcher MJ, Dick JTA, Dunn AM (2012) Diverse effects of parasites in ecosystems: linking inter-
dependent processes. Front Ecol Environ 10:186–194
Heeger T, Piatkowski U, Möller H (1992) Predation on jellyfi sh by the cephalopod Argonauta
argo . Mar Ecol Prog Ser 88:293–296
Heins DC, Birden EL, Baker JA (2010) Host mortality and variability in epizootics of
Schistocephalus solidus infecting the threespine stickleback, Gasterosteus aculeatus .
Parasitology 137:1681–1686
Hosia A, Båmstedt U (2007) Seasonal changes in the gelatinous zooplankton community and
hydromedusa abundances in Korsfjord and Fanafjord, western Norway. Mar Ecol Prog Ser
351:113–127
Houghton JDR, Doyle TK, Davenport J, Lilley MKS, Wilson RP, Hays GC (2007) Stranding
events provide indirect insights into the seasonality and persistence of jellyfi sh medusae
(Cnidaria: Scyphozoa). Hydrobiologia 589:1–13
Ishii H, Båmstedt U (1998) Food regulation of growth and maturation in a natural population of
Aurelia aurita (L.). J Plankton Res 20:805–816
Jansen PA, Kristoffersen AB, Viljugrein H, Jimenez D, Aldrin M, Stien A (2012) Sea lice as a
density-dependent constraint to salmonid farming. Proc Roy Soc Lond Ser B 279:2330–2338
K.A. Pitt et al.
101
Jarms G, Båmstedt U, Tiemann H, Martinussen MB, Fosså JH (1999) The holopelagic life cycle of
the deep-sea medusa Periphylla periphylla (Scyphozoa, Coronatae). Sarsia 84:55–65
Kikinger R (1992) Cotylorhiza tuberculata (Cnidaria: Scyphozoa) – Life history of a stationary
population. Mar Ecol 13:333–362
Lafferty KD (2008) Ecosystem consequences of fi sh parasites. J Fish Biol 73:2083–2093
Larson RJ (1986) Changes in the standing stocks, growth rates, and production rates of gelatinous
predators in Saanich Inlet, British Columbia. Mar Ecol Prog Ser 33:89–98
Lauckner G (1980) Diseases of Cnidaria. In: Kinne O (ed) Diseases of marine animals, Volume I:
general aspects, Protozoa to Gastropoda. Wiley, Chichester, pp 167–237
Laval P (1972) Comportement, parasitisme et écologie d’ Hyperia schizogeneio s Stebb.
(Amphipode Hypéride) dans le plancton de Villefranche-sur-mer. Ann de l’Instit Océanogr
Paris 48:49–74
Laval P (1980) Hyperiid amphipods as crustacean parasitoids associated with gelatinous zooplank-
ton. Oceanogr Mar Biol Annu Rev 18:11–56
Lebrato M, Jones DOB (2009) Mass deposition event of Pyrosoma atlanticum carcasses off Ivory
Coast (West Africa). Limnol Oceanogr 54:1197–1209
Lebrato M, Pahlow M, Oschlies A, Pitt KA, Jones DOB, Molinero JC, Condon RH (2011) Depth
attenuation of organic matter export associated with jelly falls. Limnol Oceanogr
56:1917–1928
Lebrato M, Pitt KA, Sweetman AK, Jones DOB, Cartes JE, Oschlies A, Condon RH, Molinero JC,
Adler L, Gaillard C, Lloris D, Billett DSM (2012) Jelly-falls historic and recent observations:
a review to drive future research directions. Hydrobiologia 690:227–245
Lebrato M, Jesus Mendes P, Steinberg DK, Cartes JE, Jones BM, Birsa LM, Benavides R, Oschiles
A (2013) Jelly biomass sinking speed reveals a fast carbon export mechanism. Limnol
Oceanogr 58:1113–1122
Lilley MKS, Houghton JDR, Hays GC (2009) Distribution, extent of inter-annual variability and diet
of the bloom-forming jellyfi sh Rhizostoma in European waters. J Mar Biol Assoc UK 89:39–48
Lilley MKS, Beggs SE, Doyle TK, Hobson VJ, Stromberg KHP, Hays GC (2011) Global patterns
of epipelagic gelatinous zooplankton biomass. Mar Biol 158:2429–2436
Lin Y-CJ, Grigoriev NG, Spencer AN (2000) Wound healing in jellyfi sh striated muscle involves
rapid switching of two modes of cell mobility and a change in the source of regulatory calcium.
Dev Biol 225:87–100
Link JS, Ford MD (2006) Widespread and persistent increase of Ctenophora in the continental
shelf ecosystem off NE USA. Mar Ecol Prog Ser 320:153–159
Lo WT, Chen IL (2008) Population succession and feeding of scyphomedusae, Aurelia aurita , in a
eutrophic tropical lagoon in Taiwan. Estuar Coast Shelf Sci 76:227–238
Lucas CH (1996) Population dynamics of Aurelia aurita (Scyphozoa) from an isolated brackish
lake, with particular reference to sexual reproduction. J Plankton Res 18:987–1007
Lucas CH, Williams JA (1994) Population dynamics of the scyphomedusa Aurelia aurita in
Southampton Water. J Plankton Res 16:879–895
Lucas CH, Hirst AG, Williams JA (1997) Plankton dynamics and Aurelia aurita production in two
contrasting ecosystems: Comparisons and consequences. Estuar Coast Shelf Sci 45:209–219
Lützen J (2005) Amphipoda (amphipods). In: Rohde K (ed) Marine parasitology. CSIRO
Publishing, Collingwood, pp 165–169
Madin LP, Harbison GR (1977) The associations of Amphipoda Hyperiidea with gelatinous
zooplankton – I. Associations with Salpidae. Deep-Sea Res 24:449–463
Malej A, Malej M (1992) Population dynamics of the jellyfi sh Pelagia noctiluca (Forsskål, 1775).
Marine eutrophication and population dynamics. In: Proceedings of 25th EMBS, pp 215–219
Marcogliese DJ (1995) The role of zooplankton in the transmission of helminth parasites to fi sh.
Rev Fish Biol Fish 5:336–371
Martin LE, Dawson MN, Bell LJ, Colin PL (2006) Marine lake ecosystem dynamics illustrate
ENSO variation in the tropical western Pacifi c. Biol Lett 2:144–147
Martorelli SR, Cremonte F (1998) A proposed three-host life history of Monascus fi liformis
(Rudolphi, 1819) (Digenea: Fellodistomidae) in the southwest Atlantic Ocean. Can J Zool
76:1198–1203
4 Bloom and Bust: Why Do Blooms of Jellyfi sh Collapse?
102
Matsakis S, Conover RH (1991) Abundance and feeding of medusae and their potential impact as
predators on other zooplankton in Bedford Basin (Nova Scotia, Canada) during Spring. Can J
Fish Aquat Sci 48:1419–1430
Mayer AG (1914) The law governing the loss of weight in starving Cassiopea . Pap Tortugas Lab
Carnegie Inst Wash 6:55–83
McDermott JJ, Zubkoff PL, Lin AL (1982) The occurrence of the anemone Peachia parasitica as
a symbiont in the scyphozoan Cyanea capillata in the lower Chesapeake Bay. Estuaries
5:319–321
Metz P (1967) On the relations between Hyperia galba Montagu (Amphipoda, Hyperiidae) and its
host Aurelia aurita in the Isefjord area (Sjaelland, Denmark). Vidensk Medd Dan Naturhist
Foren 130:85–108
Miglietta MP, Rossi M, Collin R (2008) Hydromedusa blooms and upwelling events in the Bay of
Panama, Tropical East Pacifi c. J Plankton Res 30:783–793
Mills CE (1993) Natural mortality in NE Pacifi c coastal hydromedusae: Grazing predation, wound
healing and senescence. Bull Mar Sci 53:194–203
Möller H (1980) Population dynamics of Aurelia aurita medusae in Kiel Bight, Germany (FRG).
Mar Biol 60:123–128
Møller LF, Riisgård HU (2007) Population dynamics, growth and predation impact of the common
jellyfi sh Aurelia aurita and two hydromedusae, Sarsia tubulosa and Aequorea vitrina , in
Limfjorden (Denmark). Mar Ecol Prog Ser 346:153–165
Mutlu E (2001) Distribution and abundance of moon jellyfi sh ( Aurelia aurita ) and its zooplankton
food in the Black Sea. Mar Biol 138:329–339
Ohtsuka S, Koike K, Lindsay D, Nishikawa J, Miyake H, Kawahara M, Mujiono N, Hiromi J,
Komatsu H (2009) Symbionts of marine medusae and ctenophores. Plankton Benth Res 4:1–13
Olesen NJ, Frandsen K, Riisgård HU (1994) Population dynamics, growth and energetics of
jellyfi sh Aurelia aurita in a shallow fjord. Mar Ecol Prog Ser 105:9–18
Pagès F (2000) Biological associations between barnacles and jellyfi sh with emphasis on the
ectoparasitism of Alepas pacifi ca (Lepadomorpha) on Diplulmaris malayensis (Scyphozoa).
J Nat Hist 34:2045–2056
Papathanassiou E, Panayotidis P, Anganostaki K (1987) Notes on the biology and ecology of the
jellyfi sh Aurelia aurita Lam. In Elefsis Bay (Saronikos Gulf, Greece). Mar Ecol 8:49–58
Pertsova NM, Kosobokova KN, Prudkovsky AA (2006) Population size structure, population dis-
tribution, and life cycle of the Hydromedusa Aglantha digitale (O.F. Müller, 1766) in the White
Sea. Oceanology 46:228–237
Pitt KA, Kingsford MJ (2000) Geographic separation of stocks of the edible jellyfi sh Catostylus
mosaicus (Rhizostomeae) in New South Wales, Australia. Mar Ecol Prog Ser 196:143–155
Pitt KA, Clement AL, Connolly RM, Thibault-Botha D (2008) Predation by jellyfi sh on large and
emergent zooplankton: implications for benthic-pelagic coupling. Estuar Coast Shelf Sci
76:827–833
Pitt KA, Welsh DT, Condon RH (2009a) Infl uence of jellyfi sh blooms on carbon, nitrogen and
phosphorus cycling and plankton production. Hydrobiologia 616:133–149
Pitt KA, Connolly RM, Meziane T (2009b) Stable isotope and fatty acid tracers in energy and
nutrient studies of jellyfi sh: a review. Hydrobiologia 616:119–132
Pitt KA, Duarte CM, Lucas CH, Sutherland KR, Condon RH, Mianzan H, Purcell JE, Robinson
KL, Uye S-I (2013) Jellyfi sh body plans provide allometric advantages beyond low carbon
content. PLoS One 8(8):e72683
Primo AL, Marques SC, Falcão J, Crespo D, Pardal MA, Azeiteiro U (2012) Environmental forcing
on jellyfi sh communities in a small temperate estuary. Mar Environ Res 79:152–159
Purcell JE, Arai MN (2001) Interactions of pelagic cnidarians and ctenophores with fi sh: a review.
Hydrobiologia 451:27–44
Purcell JE, Decker MB (2005) Effects of climate on relative predation pressure by scyphomedusae
and copepods in Chesapeake Bay during 1987–2000. Limnol Oceanogr 50:376–387
Purcell JE, White JR, Nemazie DA, Wright DA (1999) Temperature, salinity and food effects on
asexual reproduction and abundance of the scyphozoan Chrysaora quinquecirrha . Mar Ecol
Prog Ser 180:187–196
K.A. Pitt et al.
103
Reimann L, Titelman J, Båmstedt U (2006) Links between jellyfi sh and microbes in a jellyfi sh
dominated fjord. Mar Ecol Prog Ser 325:29–42
Reitzel AM, Sullivan JC, Finnerty JR (2006) Qualitative shift to indirect development in the
parasitic sea anemone Edwardsiella lineata . Integr Comp Biol 46:827–837
Reitzel AM, Sullivan JC, Brown BK, Chin DW, Cira EK, Edquist SK, Genco BM, Joseph OC,
Kaufman CA, Kovitvongsa K, Munoz MM, Negri TL, Taffel JR, Zuehike RT, Finnerty JR
(2007) Ecological and developmental dynamics of a host-parasite system involving a sea
anemone and two ctenophores. J Parasitol 93:1392–1402
Rhode SC, Pawlowski M, Tollrian R (2001) The impact of ultraviolet radiation on the vertical
distribution of zooplankton of the genus Daphnia . Nature 412:69–72
Riisgård HU, Barth-Jensen C, Madsen CV (2010) High abundance of the jellyfi sh Aurelia aurita
excludes the invasive ctenophore Mnemiopsis leidyi to establish in a shallow cove (Kertinge
Nor, Denmark). Aquat Invasions 5:347–356
Rippingale RJ, Kelly SJ (1995) Reproduction and survival of Phyllorhiza punctata (Cnidaria:
Rhizostomeae) in a seasonally fl uctuating salinity regime in Western Australia. Mar Freshwater
Res 46:1145–1151
Salonen K, Högmander P, Langenberg V, Mölsä H, Sarvala J, Tarvainen A, Tiirola M (2012)
Limnocnida tanganyicae medusae (Cnidaria: Hydrozoa): a semiautonomous microcosm in the
food web of Lake Tanganyika. Hydrobiologia 690:97–112
Schneider G, Behrends G (1994) Population dynamics and the trophic role of Aurelia aurita
medusae in the Kiel Bight and western Baltic. ICES J Mar Sci 51:359–367
Selander E, Møller L, Sundberg P, Tiselius P (2010) Parasitic anemone infects the invasive
ctenophore Mnemiopsis leidyi in the North East Atlantic. Biol Invasions 12:1003–1009
Sexton MA, Hood RR, Sarkodee-adoo J, Liss AM (2010) Response of Chrysaora quinquecirrha
medusae to low temperature. Hydrobiologia 645:125–133
Spangenberg DB (1965) Cultivation of the life stages of Aurelia aurita under controlled conditions.
J Exp Zool 159:303–318
Spaulding JG (1972) The life cycle of Peachia quinquecapitata , an anemone parasitic on medusae
during its larval development. Biol Bull 143:440–453
Sun S, Lin Y, Sun X (2012) Changes in the small-jellyfi sh community in recent decades in Jiaozhou
Bay, China. Chin J Oceanol Limnol 30:507–513
Svendsen YS (1990) Hosts of third stage larvae of Hysterothylacium sp. (Nematoda, Anisakidae)
in zooplankton from outer Oslofjord, Norway. Sarsia 75:161–167
Tinta T, Kogovšek T, Malej A, Turk V (2012) Jellyfi sh modulate bacterial dynamic and community
structure. PLoS One 7:e39274
Towanda T, Thuesen EV (2006) Ectosymbiotic behavior of Cancer gracilis and its trophic rela-
tionships with its host Phacellophora camtschatica and the parasitoid Hyperia medusarum .
Mar Ecol Prog Ser 315:221–236
Vannuci-Mendes M (1944) Sobre a larva de Dibothriorhynchus dinoi , sp. n. parasita dos
Rhizostomata. Arquivos do Museu Paranaense 4:47–81
Williams R, Robins D (1981) Seasonal variability in abundance and vertical-distribution of
Parathemisto gaudichaudi (Amphipoda: Hyperiidea) in the North East Atlantic Ocean. Mar
Ecol Prog Ser 4:289–298
Wintzer AP, Meek MH, Moyle PB (2011) Life history and population dynamics of Moerisia sp., a
non-native hydrozoan, in the upper San Francisco Estuary (U.S.A). Estuar Coast Shelf Sci
94:48–55
Yamamoto J, Hirose M, Ohtani T, Sugimoto K, Hirase K, Shimamoto N, Shimura T, Honda N,
Fujimori Y, Mukai T (2008) Transportation of organic matter to the sea fl
oor by carrion
falls of the giant jellyfi sh Nemopilema nomurai in the Sea of Japan. Mar Biol 153:
311–317
Yip SY (1984) Parasites of Pleurobrachia pileus Muller, 1776 (Ctenophora), from Galway Bay,
western Ireland. J Plankton Res 6:107–121
Zahn M (1981) Wie alt können Scyphomedusen warden? Zoologische Beitraege 27:491–495
Zeleney C (1907) The effect of degree of injury, successive injury and functional activity upon
regeneration in the scyphomedusan, Cassiopea xamachana . J Exp Zool 5:265–274
4 Bloom and Bust: Why Do Blooms of Jellyfi sh Collapse?
