Persistence, impacts and environmental drivers of covert infections in invertebrate hosts
Fontes et al. Parasites & Vectors
Persistence, impacts and environmental drivers of covert infections in invertebrate hosts
Inês Fontes 0 2
Hanna Hartikainen 1 4
Chris Williams 3
Beth Okamura 0
0 Department of Life Sciences , Natural History Museum, Cromwell Road, London SW7 5BD , UK
1 EAWAG, Department of Aquatic Ecology , Überlandstrasse 133, CH-8600 Dübendorf , Switzerland
2 Scottish Fish Immunology Research Centre, Aberdeen University , Aberdeen AB24 2TZ , UK
3 Environment Agency, National Fisheries Laboratory , Brampton, Cambridgeshire PE28 4NE , UK
4 ETH Zürich, Institute of Integrative Biology (IBZ) , Zürich , Switzerland
Background: Persistent covert infections of the myxozoan, Tetracapsuloides bryosalmonae, in primary invertebrate hosts (the freshwater bryozoan, Fredericella sultana) have been proposed to represent a reservoir for proliferative kidney disease in secondary fish hosts. However, we have limited understanding of how covert infections persist and vary in bryozoan populations over time and space and how they may impact these populations. In addition, previous studies have likely underestimated covert infection prevalence. To improve our understanding of the dynamics, impacts and implications of covert infections we employed a highly sensitive polymerase chain reaction (PCR) assay and undertook the first investigation of covert infections in the field over an annual period by sampling bryozoans every 45 days from three populations within each of three rivers. Results: Covert infections persisted throughout the year and prevalence varied within and between rivers, but were often > 50%. Variation in temperature and water chemistry were linked with changes in prevalence in a manner consistent with the maintenance of covert infections during periods of low productivity and thus poor growth conditions for both bryozoans and T. bryosalmonae. The presence and increased severity of covert infections reduced host growth but only when bryozoans were also investing in the production of overwintering propagules (statoblasts). However, because statoblast production is transitory, this effect is unlikely to greatly impact the capacity of bryozoan populations to act as persistent sources of infections and hence potential disease outbreaks in farmed and wild fish populations. Conclusions: We demonstrate that covert infections are widespread and persist over space and time in bryozoan populations. To our knowledge, this is the first long-term study of covert infections in a field setting. Review of the results of this and previous studies enables us to identify key questions related to the ecology and evolution of covert infection strategies and associated host-parasite interactions.
Tetracapsuloides bryosalmonae; Fredericella sultana; Proliferative kidney disease; Myxozoans; Bryozoans; Productivity; Temperature; Host condition-dependent effects; Covert infection strategies; Fish disease reservoirs
Covert or latent infections are non-infective, persistent
forms of a parasitic infection that are asymptomatic in
host populations [
]. Stages causing covert infections
may be latent or may undergo low levels of replication
]. When activated, covert infections become overt and
result in detectable diseases [
] that may be lethal [
Covert infections are typically caused by agents whose
small sizes enable persistence as cryptic stages within
much larger hosts without invoking obvious harm.
Covert viral infections have received most attention, for
example those causing human immunodeficiency virus
] and hepatitis C [
] in humans, and those
associated with insect diseases, such as granuloviruses (GV)
], baculoviruses [
] and nucleopolyhedroviruses [
Bacteria can also cause covert infections in many
organisms, including humans (e.g. tuberculosis), fish [
]. Eukaryotic organisms causing covert
infection include the microsporidian, Encephalitozoon
], and protozoans, such as Plasmodium
]. At present, however, we are aware of
only a single group of metazoans that demonstrate
covert infection dynamics - myxozoans that are
associated with freshwater bryozoan [
] and annelid [
hosts. Myxozoans are a clade of endoparasitic cnidarians
with complex life-cycles, exploiting invertebrates and
vertebrates as primary and secondary hosts, respectively.
Morphological simplification and miniaturisation, along
with a capacity for multiplication within hosts, have
enabled myxozoans uniquely to converge on infection
strategies associated with microparasites [
infections of malacosporean myxozoans in freshwater
bryozoans (benthic, colonial invertebrates) have been
examined in two systems: Tetracapsuloides
bryosalmonae in the bryozoan Fredericella sultana [
Buddenbrockia allmani in the bryozoan Lophopus
]. The former system has been
particularly investigated as T. bryosalmonae is the causative
agent of proliferative kidney disease (PKD) of salmonid
fish. It has been argued that covert infections that cause
little to no adverse effects enable long term persistence
of T. bryosalmonae in bryozoan populations and hence a
disease reservoir for fish [
]. Confirmation that
infected overwintering stages of bryozoans (statoblasts)
are viable [
] reveals one mechanism for such
infection persistence in bryozoan populations. In addition,
transmission of T. bryosalmonae to fish upon exposure
to colonies deriving from infected statoblasts [
indicates that infection of statoblasts creates an effective
disease reservoir. However, observations and collections of
live colonies of F. sultana during winter periods [
suggest that covert infections might persist in bryozoan
colonies throughout the year. If so, overwintering
colonies may offer an additional and direct means of
infection persistence within populations and this could also
contribute to the disease reservoir for fish.
Molecular diagnostics have shown that T. bryosalmonae
infections persist over weeks to months at varying
prevalences in bryozoan populations. One study found
prevalences to range from 0 to 53% during early April to
mid-September in three bryozoan populations situated
along a short (~15 m) stretch of the River Cerne (Dorset,
]. A second study revealed infection prevalences
of 0–17% in bryozoans transplanted into a single locality
on the River Itchen (Hampshire, UK) during late June to
early August [
]. Neither study distinguished covert from
overt infections. How covert infection prevalences vary
over a full annual cycle in bryozoan populations remains
unknown, and variation in prevalences in bryozoan
populations within and between rivers is poorly understood.
Furthermore, previous studies used polymerase chain
reaction (PCR) primers that are likely to have
underestimated infection prevalences due to lower sensitivity than
that of more recently designed primers.
To improve our understanding of the dynamics,
impacts and implications of covert infections in
bryozoan populations we utilised a recently developed and
highly sensitive PCR assay to address the following
hypotheses: (i) Bryozoan hosts harbour covert infections
of T. bryosalmonae throughout the year, contributing to
the disease reservoir for fish; (ii) Covert infection
prevalences and severities vary within and between rivers; (iii)
Covert infection prevalences and severities are
influenced by environmental conditions; and (iv) Covert
infections have no impact on bryozoans in the field, a
prediction in keeping with limited evidence of little to
no effect on host growth and statoblast production in
laboratory studies (see below).
Freshwater bryozoans are colonial, suspension feeding
invertebrates that are ubiquitous but overlooked
residents of freshwater environments. In temperate
regions, colonies grow during warmer months of the
year by budding new zooids, each of which has a
tentacular crown (the lophophore) used for suspension
]. The most common bryozoan host of T.
bryosalmonae is F. sultana, which can form dense stands
of colonies in the interstices of submerged roots of
riparian trees [
]. Fredericella sultana grows as branching,
tubular colonies and reproduces mainly asexually by
colony fragmentation and the production of
overwintering, dormant seed-like stages called statoblasts
]. Statoblasts enclose germinal tissues and hatch to
give rise to small colonies in spring as temperatures
increase. F. sultana can also persist during winter as live
Tetracapsuloides bryosalmonae cycles between two
developmental stages within bryozoans, resulting in
covert and overt infections. The former are characterised by
cryptic stages consisting of single cells associated with
the body wall [
] and are detectable by PCR. Controlled
laboratory studies provide evidence that covert
infections are largely avirulent. Thus, for bryozoans collected
from one river system (the River Cerne [
]) there were
no detectable effects of covert infection on host growth,
except when hosts were also investing in statoblast
production, nor in the propensity to produce statoblasts. In
addition, the number of statoblasts produced was not
impacted by whether colonies were covertly infected in
bryozoans originating from two other river systems (the
Rivers Avon and Dun [
]). Furthermore, infected
statoblasts from three river systems (the Rivers Itchen, Dun
and Lyssbach) have been shown to be viable [
to exhibit greater hatching success than uninfected
statoblasts . Covert infections are also present in
colony fragments that detach from parental colonies in
the field [
]. Both statoblast production and colony
fragmentation are modes of host propagation that T.
bryosalmonae can exploit to effectively achieve vertical
transmission to new bryozoan colonies. Overt infections
involve the development of multicellular sacs that are
readily observed with a stereomicroscope circulating
freely within the body cavity (e.g. [
19, 28, 30
mature within sacs and are released into the water to
infect fish. Overt infections are virulent, impacting
bryozoan growth [
] and causing temporary castration by
inhibiting statoblast production [
]. Overt infection
prevalences are high in late spring and autumn [
when the hosts are in good condition [
] and able
to support the development of these relatively large,
rapidly growing and energetically costly stages [
Sac production in highly stressed, food-deprived
bryozoans  demonstrates that overt infections can
also develop in response to potential host death, a form
of terminal investment. This response may explain
infection of fish in winter [
] when conditions for bryozoans
Study sites and sample collection
Fredericella sultana colonies were collected every 45 days
over 1 year from three rivers in southern England: the
River Avon (near Bickton, Hampshire), the River Dun
(Hungerford, Berkshire) and the River Itchen (Winchester,
Hampshire). The Rivers Avon and Dun were sampled for
12 months beginning in October 2012, staggering the
dates between sites to accommodate processing. The River
Itchen was sampled for 12 months beginning in October
2013. The sites were chosen based on the presence of
PKD on fish farms associated with the rivers (O.
Robinson, pers. comm.) and the presence of F. sultana
populations on submerged tree root systems (B. Okamura,
pers. obs.). Up to 100 branches of bryozoan colonies
(depending on availability) were collected haphazardly
using forceps from each of three tree root systems in each
river. Additional file 1: Table S1 provides locations of the
tree root systems sampled in each river. Branches
detached from colonies were placed into individual 15 ml
plastic tubes filled with river water and were kept at 4 °C
for 24 h until dissection (see below).
Temperature, flow speed and dissolved oxygen (DO) were
recorded on almost every sampling trip. Temperature and
DO were measured in each river at the root located furthest
downstream using an oximeter (WTW Oxi330). To
characterise incoming flow experienced by the root system, flow
speed was measured in a position directly next to the initial
development of the root system and at one third of the
river depth from the surface using an open channel
electromagnetic flow meter (Valeport 8008/801) and positioning
the flat flow sensor in an upstream direction to estimate
mean speed of flow (over 60 s). Flow was recorded at the
same position and equivalent depths during each sampling
trip. During the last four sampling trips (i.e. from April
onwards) samples were taken for water chemistry,
productivity, turbidity and conductivity analyses
(Additional file 2: Table S2) next to the root located furthest
downstream in each river (except in the River Itchen where
samples were taken from root 2) according to standard
procedures. The water samples were stored at 4 °C and
analysed within 24 h by the Environment Agency’s National
Laboratory Service (NLS), Starcross, UK.
Colony attributes and infection state
Detached branches (henceforth referred to as colonies)
were observed using a stereomicroscope to determine
their status, i.e. whether they were alive or dead (no
living zooids) and in both cases, whether they contained
statoblasts. The sizes of a subset of live colonies
(numbers counted according to time permitting and ranging
from 41 to 87 colonies per root system) were
determined by counting the total number of live zooids
(lophophores and digestive tracts present). All colonies were
then dissected (using dissection tools sterilised with
ethanol) to determine whether mature statoblasts and
overt infections (sacs) were present. Statoblasts were
considered to be mature if they had assumed the typical
brown colouration due to tanning of chitin. The sizes of
dead colonies that contained mature statoblasts were
recorded as zero. Following dissection, colonies were
preserved in 100% ethanol and stored at -20 °C (with the
exception of dead colonies). Data on colony size
(excluding dead colonies) and statoblast production were
used to analyse the impacts of covert infection on
bryozoans in each river system.
Colonies that were not overtly-infected were screened
using PCR to characterise proportions of uninfected and
covertly-infected colonies. The number of colonies
screened by PCR was determined by availability or, when
sufficiently numerous, by haphazardly choosing subsets
of colonies from each root (up to 55 colonies per root).
DNA was then extracted from each bryozoan colony (i.e.
the tissue present in detached branches) using a
modified hexadecyltrimethylammonium bromide (CTAB)
protocol (see Additional file 3). Covert infections were
detected using a PCR assay with cycling conditions as in
Hartikainen et al. [
] based on primers diagnostic for
T. bryosalmonae 18S rDNA [514F_new (5′-ATT CAG
GTC CAT TCG TGA GTA ACA AGC-3′) and 776R
(5′-CTG CCC TTA ATT GGG TGT ATC AGC-3′)] to
produce an amplicon of 244 bp. This set of primers
appears to be highly sensitive to T. bryosalmonae,
indicating much greater infection prevalences in
bryozoan populations than those revealed by different
primers used in previous studies [
]. The PCR products
(3 μl) were run on a 1.5% agarose electrophoretic gel.
Infection intensity was characterised by the PCR
product’s molecular weight (ng/μl) estimated by the gel band
intensity of two individual replicate PCRs [
]. This is a
semi-quantitative method of estimating colony infection
intensities (precluding accurate estimates of very strong/
weak signals [
]). In view of this potential constraint
we amplified and gel-analysed all samples in duplicate
and prepared, ran, photographed and analysed gels using
identical conditions [
]. Infection severity was then
calculated by dividing infection intensity by colony size.
One sample that was positive for T. bryosalmonae for
each root in each river was selected randomly (total of 9
samples) and verified by direct sequencing using an ABI
PRISM® 3700xl DNA analyser (Applied Biosystems,
Foster City, USA) and BigDye v1.1 chemistry. Bryozoans
were retained for 24 h prior to dissection which allowed
voiding faeces (and hence ingested spores) and
promoted breakdown of any T. bryosalmonae spores
retained on surfaces or in the water (spores degrade
between 12 and 24 h, as shown in [
]). Dishes were
rinsed with warm water and dissection equipment with
ethanol between colonies. We also employed negative
controls in DNA extractions and in PCR runs.
All statistical analyses were conducted using R version
]. Principal components analysis (PCA) was
performed on the environmental variables,
circumventing the problem of multi-collinearity, to create
uncorrelated axes and to investigate variation associated with
rivers by cluster analysis. PCA was run on centred and
scaled variables (i.e. mean = 0 and standard deviation = 1)
and calculated from the correlation matrix using
singular value decomposition (SVD) with the stats package
version 3.1.1 [
]. PC biplots and ellipses with normal
probability contours set to 68% were created using the
ggbiplot package version 0.55 [
]. A Scree plot was used
to choose the number of principal components (PCs) or
axes to consider as meaningful representations of the
data and to include as fixed explanatory variables in
subsequent Generalised Linear Mixed Models (GLMMs).
Differences in live colony size were compared across
sampling trips using a Generalised Linear Model (GLM)
with a Quasi-Poisson error distribution. The proportion
of colonies producing statoblasts (both live and dead)
was compared amongst rivers using a GLM with a
binomial error distribution.
Relationships between infection state and explanatory
variables, including environmental variables and colony
attributes, were assessed with GLMMs following the
methods described in Zuur et al. [
measures of colonies used as response variables included
covert infection status (present or absent) and,
conditional on covert infection being present, the infection
severity. We used the package lme4 version 1.1-7 [
assuming a binomial error distribution to analyse
infection status, and the package nlme version 3.1-117 [
analyse infection severity. Variability in the response
variable amongst rivers, roots and sampling trips was
investigated graphically. Random effects models were
then built to determine the optimal random structure
using restricted maximum likelihood estimation (REML)
and maximum likelihood estimation (ML) for infection
severity and status, respectively. Univariable analyses
were used to explore the influence of each fixed
explanatory variable (Table 1). Only those variables with
Pvalues < 0.25 were included in a maximal model using
ML estimation following a visual inspection of the data.
Variables and interactions were then removed from the
maximal model in a stepwise fashion by establishing
whether their removal caused a significant change in the
model’s Akaike Information Criterion (AIC) value.
In view of our broad range of results the associated
statistical tests and their inferences are incorporated
collectively in table format to enable ready comparison
Temperatures in the three rivers generally declined from
October to December and remained low until early spring.
Temperatures in the River Dun did not vary as much as in
the other two rivers (Additional file 4: Figure S1). The
results for all environmental variables (except
temperature) and for flow speed are provided in
Additional file 2: Table S2 and Additional file 5: Table S3,
respectively. Data for 21 environmental variables,
including temperature and flow speed, measured for the last four
sampling trips to the Rivers Avon and Dun were suitable
for PCA (the large amount of missing data for the River
Itchen precluded its use). Two PCs explained 81% of the
variance in the data - PC1 which represents the two rivers
explained some 56% of the variance and PC2 which
represents the four sampling trips explained some 25% of
the variance (see Additional file 6: Figure S2 and
Additional file 7: Table S4). PC1 had strong positive
loadings for Bacterial Oxygen Demand (BOD), nitrogen
associated variables, chloride, orthophosphate, magnesium,
coliforms (presumptive and confirmed) and temperature,
and strong negative loadings for DO and alkalinity. PC2
had strong positive loadings for chlorophyll-a and
turbidity, and strong negative loadings for temperature,
alkalinity, orthophosphate, conductivity, hardness and calcium.
Ellipses and colour coded data demonstrate that rivers
and sampling trips are not overlapping in the PC space
(Additional file 6: Figure S2). These results show that
rivers differ in water chemistry and that this varies less
between sampling trips 6–8 on the River Avon.
Bryozoan hosts over time and space
Up to 300 F. sultana colonies were collected during each
of eight sampling trips from the Rivers Avon (total
number of colonies 1563), Dun (total number of colonies
1562) and Itchen (total number of colonies 1337)
(Table 2, Fig. 1a). The majority of the colonies collected
on each sampling date were alive but the number of
dead colonies increased during the winter, especially in
the Rivers Dun and Itchen. Collecting material generally
became more difficult with the onset of winter when
many bryozoan colonies degenerated. Live colonies had
disappeared completely in some cases by late winter/
early spring (on roots in the River Avon and from some
roots in the River Itchen) (Table 2). However new
growth enabled collection of live colonies in large
numbers in spring and summer from all roots in each river
(Fig. 1a, Table 2). Accordingly, the mean sizes of live
colonies significantly varied over time in all three rivers
and were generally smallest in winter (Fig. 1b, Table 3:
Test A1). The proportion of colonies with statoblasts
was significantly different amongst rivers (Table 3: Test
A2). Statoblasts were present in a minority of colonies in
the Rivers Avon and Dun and were most commonly
observed in late summer and autumn. Statoblasts were
almost entirely absent from bryozoans collected from the
River Itchen (Table 2).
Tetracapsuloides bryosalmonae infections over time and
Covert infections were present throughout the year in all
three rivers (Fig. 2a). GLM indicated that the proportion
of covert infections varied significantly amongst rivers
(Table 3: Test B1) and for sampling trips within each
river (Table 3: Test B2). In the River Avon the mean
prevalence of covert infections was lowest in October
(23%) and highest in January (76%). In the River Dun,
mean covert infection prevalences were lowest in April
(36%) and, as in the Avon, highest in January (92%). In
contrast, in the River Itchen mean covert infection
prevalences were lowest in January (33%) and highest in
October (73%). Covert infection prevalences were also
variable amongst roots within rivers (ranging between 3
and 100% at any given time (Table 2, Table 3: Test B3).
Overt infections were rare (detected in 0–5% of
colonies per root at a given time) (Table 2) but were
present in at least one river in most sampling periods,
apart from during January-March, when they were not
encountered. The very low prevalences of overt infection
precluded any analyses of infection patterns. There is a
suggestion that the numbers of sacs increased in late
summer in all rivers (Table 2).
Covert infections in relation to host and environmental conditions
Rivers, roots nested within rivers and sampling trips
nested within roots explained a large amount of
variation in covert infection status. These variables were
therefore included as random effects within subsequent
mixed models to assess the significance of host
characters on covert infection status (Table 1). The two-way
interaction between colony size and statoblast presence
had a significant effect on infection status (Table 3: Test
B4), with a unit increase in size of colonies containing
statoblasts decreasing the likelihood of colonies being
covertly-infected by 0.920 times (odds ratio). This result
suggests that the utilisation of resources by concomitant
covert infections reduces host growth if bryozoans are
also investing in statoblast production. Notably there
were no other discernible relationships between covert
infection and colony size in general nor in the
propensity to produce statoblasts.
Rivers, roots nested within rivers and sampling trips
nested within roots were also included as random effects
in mixed models to assess the significance of
environmental variables (summarised by PC1 and PC2 values) on
covert infection status in the Rivers Avon and Dun. The
three-way interaction between colony size, statoblast
presence and PC1 had a significant effect on covert
infection status (Table 3: Test B5). An increase in PC1’s
positive loadings (mainly productivity and temperature) and a
decrease in PC1’s negative loadings (e.g. DO), were
associated with an increase in the size of statoblast-producing
colonies - in turn this was associated with
statoblastbearing colonies being 0.953 times (odds ratio) less likely
to be covertly infected. This pattern mirrors the above
results, suggesting an energetic drain caused by covert
infections that reduces host growth when bryozoans are
also investing in statoblast production.
Covert infection intensity values are provided in
Additional file 8: Table S5. The associated infection
severities varied over time both within and amongst rivers
(Fig. 2b). Mean covert infection severities were enhanced in
overwintering colonies in the Rivers Avon and Dun but
were highest in April in the River Itchen. Mean infection
severity was generally lowest in the River Avon. None of
the explanatory variables (Table 1) significantly affected
infection severity in mixed models with rivers, roots nested
within rivers and sampling trips nested within roots as
random effects. However, PC1 had a significant effect on
infection severity (Table 3: Test B6), with a unit increase in PC1
causing a decrease in infection severity of -0.237 (± 0.130
SE). In other words, an increase in PC1’s positive loadings
(mainly productivity and temperature) and a decrease in
PC1’s negative loadings (e.g. DO) decreased covert infection
severity. Covert infection severity (analysing data pooled
across space and time) was negatively influenced by
whether colonies were producing statoblasts, with
statoblast presence associated with a -0.206 (± 0.076 SE)
decrease in infection severity (Table 3: Test B7). Covert
infection intensity was similarly likely to be lower in
colonies that were producing statoblasts (Table 3: Test B8)
and was not affected by colony size (Table 3: Test B9).
Patterns of infection in space and time
Relatively few studies have assessed covert infection
dynamics in invertebrate populations [
] and most
Test Statistical test
A1 GLM (Quasi-Poisson)
aThe test statistics for Chi-square tests and GLMMs with a binomial error distribution are χ2 values
have examined prevalences of viruses in insect,
mollusc and crustacean populations over short time
periods or at single times in different years (e.g. [
]). Our programme of systematic sampling
provides evidence that bryozoan colonies are variously
present throughout the year and that covert infections
of myxozoans are continuously harboured in these
bryozoan populations. Sampling populations every
45 days revealed that covert infection prevalences
varied between the three rivers, over time within the
rivers, and amongst the three populations sampled
within each river. Bryozoan colonies became notably
harder to collect over the winter period as they
shrank in size, but collecting material every 45 days
from the same population is likely also to have
diminished populations on tree root systems
sufficiently to preclude collection in some cases. Thus, we
failed to collect bryozoans on one date in late winter
from the River Avon and colonies were absent from
two tree root systems in the River Itchen in late
February and early April. This effect of course would
not pertain to populations on other tree roots. As
spring commenced colonies became more plentiful
and populations collected from all tree roots once
again harboured covert infections. It should be noted
that adherent statoblasts were likely to be present
locally even when colonies were unavailable for
collection. Our previous research has demonstrated
that substantial proportions (between 14 and 100% in
the River Avon and between 33 and 100% in the
River Dun; calculated from data in Abd-Elfattah et al.
]) of statoblasts carry covert infections and that
moderate levels of infections can promote hatching
]. The recruitment of a considerable number of
infected progeny should therefore ensue when
seasonal conditions improve. Indeed, such recruitment
is likely to have contributed to the flush of growth
that enabled the collection of colonies from all roots
Evidence for temporal variation in T. bryosalmonae
infections in F. sultana colonies has also been obtained
in the River Cerne (Dorset) by sampling every 3 weeks
from April to September [
]. There infection
prevalences detected by PCR ranged between 0 and 53% in
bryozoan populations on three tree root systems, with
highest values occurring from April to early June
followed by declines in late summer. The PCR study did
not distinguish covert from overt infections, however
the proportions of bryozoans harbouring overt infections
(presence of sacs) was also determined independently.
Overt infection prevalences followed a similar temporal
trend (but were delayed by one sampling period) and
ranged between 0 and 41%. It can be assumed that
covert infections will have contributed to the general
prevalences detected by PCR. The levels of covert
infections reported here are generally much higher and
consistent with prevalences reported for bryozoan
populations in the Rivers Avon and Dun that were
characterised at different periods by Abd-Elfattah et al. [
The disparity in covert infection prevalences in the River
Cerne study likely reflect the use of more sensitive
primers here and in the study by Abd-Elfattah et al. [
Influence of environmental conditions
Modelling suggests that covert infection strategies may
arise if there is seasonal variation in transmission rate or
variation in host density [
]. Our results provide some
indirect evidence for both of these scenarios. GLMM
analyses revealed that covert infections are more likely
to occur and to be more severe when BOD, nitrogen,
temperature, chloride, orthophosphate, magnesium and
coliforms are lower and DO and alkalinity are higher.
Although data for environmental conditions were only
analysed for the last four sampling periods (beginning in
April) in the Rivers Avon and Dun, these results are
consistent with the maintenance of covert infections
during periods of low productivity and thus poor
conditions for both colony growth and the development of
overt infections of T. bryosalmonae as a result of low
host food availability and low temperatures. Accordingly,
covert infections were notably common and widespread
in winter in the Rivers Avon and Dun when bryozoan
density was low.
Our laboratory studies have demonstrated that the
growth of F. sultana and the development of overt
infections of T. bryosalmonae are both stimulated by
increasing temperatures [
] and food . It is therefore
possible that conditions of higher temperatures and
productivity may have promoted overt infection
development in late spring/early summer as observed in
populations in the River Cerne [
] along with the
concomitant clearance of infection in a proportion of
colonies (e.g. in some 37% of bryozoans from the River
]), resulting in a reduced likelihood of covert
infections. Although we did not detect overt infections
in any appreciable number of colonies, the period of
peak overt infection prevalences is relatively short
(weeks) and may have been missed by our 45-day
sampling regime. Other explanations for the low
detection of overt infections include patchy distributions of
infections within tree root systems and bryozoan
resistance to overt infection development.
Effects of covert infection
Our field study revealed that larger colonies that were
producing statoblasts were less likely to be covertly
infected, an effect that does not change with the
inclusion of water chemistry in the statistical models. This
may indicate that covert infections compromise growth
of colonies when they are also investing in statoblast
production. Notably, an earlier laboratory study provides
support for this scenario. Tops et al. [
] found that
growth rates of uninfected statoblast-producing colonies
were higher than those of covertly infected
statoblastproducing colonies. Their analysis was performed on
data collected from bryozoans deriving from a single site
over a limited time period and infection times and doses
were not controlled for. Nevertheless, the combined
laboratory results of Tops et al. [
] and our current
field study provide relatively strong evidence for impacts
of covert infections on growth when bryozoans are also
producing statoblasts. An alternative explanation for the
results from our field study is that large
statoblastproducing colonies are particularly healthy and resist
infection better than smaller colonies. Separating these
alternative hypotheses would require controlled
As argued above, in general, covert infections appear to
have no impact on colony growth, apart from when
colonies are investing in statoblast production. Similarly,
the ability to produce statoblasts is apparently not
compromised by covert infections. Notably, the vast majority
of colonies sampled were lacking statoblasts over the
course of our study (a total of 3886 colonies lacked
statoblasts compared to a total of 538 colonies with statoblasts;
see Alive (-stats) and Alive (+stats) in Table 2). This
suggests that the negative effects of covert infections may be
relatively rarely experienced by bryozoans at any one time
in populations. Furthermore, covert infections have been
shown to improve statoblast hatching [
], a finding that
may partly result from a trade-off between statoblast
hatching and maternal colony growth. Clearly, however, in
order to fully understand the overall effects of covert
infections and associated host-parasite interactions,
detailed and controlled studies across the bryozoan
lifecycle are required. These should include exposing
bryozoans from a range of sites to variation in environmental
conditions and characterising growth, statoblast and larval
development, recruitment from these propagules, and the
fate and contributions of colony fragments.
Covert infections, parasite persistence and disease reservoirs
The ability of T. bryosalmonae to regress or remain as
non-virulent covert infections during sub-optimal
conditions for bryozoan hosts, coupled with vertical
transmission to asexually produced offspring (statoblasts) [
and to colony fragments [
], provide a collective
means of both ensuring and amplifying persistent
infections in clonal bryozoan hosts. We demonstrate here
that covert infections are often supported by more than
50% of the bryozoan colonies in local populations. In
addition, our results imply that a relatively small
proportion (some 14% overall) of the covertly-infected
bryozoan population experiences negative effects of covert
infection, i.e. reduced growth when simultaneously
investing in statoblast production, and we note that
producing statoblasts is a transitory state. Furthermore,
results from another study  suggest that this negative
effect may be traded off against enhanced statoblast
hatching. Bryozoan populations therefore appear to
represent persistent disease-agent reservoirs for salmonid
fish. Given the appropriate environmental cues, these
covert infections can be expected to convert to overt
infections, enabling horizontal transmission to local
farmed and wild fish. The presence, persistence and
impacts of this disease reservoir are demonstrated by
recurrent annual outbreaks of PKD in which up to 100%
of farmed fish can be infected [
] and the potential for
serious disease problems in wild fish populations [
Comparative biology of covert infections in invertebrate hosts
Collective research suggests that malacosporeans
commonly employ covert infection strategies in bryozoan
hosts. The repeated appearance and disappearance of sacs
of T. bryosalmonae in colonies of F. sultana [
18, 28, 45
and of Buddenbrockia allmani in colonies of Lophopus
] in laboratory culture provide graphic
demonstration of the retention of cryptic stages during
covert infection. It is clear that covert infections can be
widespread and commonly reach prevalences exceeding
50% in populations of both F. sultana and L. crystallinus
]; data reported here). Covert infection prevalences of
malacosporeans forming vermiform stages in bryozoan
populations remain unknown. The high infection
prevalences of sac-forming malacosporeans are explained
by transmission of infection by malacospores released
from intermediate fish hosts and by ‘vertical’ transmission
of covert infections to colony fragments [
]. We know relatively little about
transmission from fish although pilot studies demonstrate that this
is achieved over a relatively brief period of time to a
minority of colonies, i.e. ~15% (6 of 41 colonies) and ~8%
(1 of 12 colonies) for the bryozoans, F. sultana and
Plumatella fungosa over 4 and 2 week periods,
respectively (Table 2) [
]. Substantial contributions from vertical
transmission are therefore implicated and these should
facilitate both the persistence and amplification of
infection in local populations. There is a body of evidence
for transport of statoblasts by waterfowl (e.g. [
Because covert infections are present in many statoblasts,
can enhance statoblast hatching [
] and result in overt
infections that are transmissive to fish [
], such transport
is likely to introduce malacosporean infections to new sites.
Infection of a larva of F. sultana by T. bryosalmonae has
been demonstrated by PCR (Fontes unpub. data). However,
the rarity of larval production in some F. sultana
populations suggests this form of transmission makes a minor
contribution to malacosporean infection persistence.
There is little evidence that covert infections of T.
bryosalmonae are the result of host suppression of overt
infection development [
]. Hosts experiencing poor
conditions (such as low food levels) are often
], yet covert infections are particularly linked with
such unproductive conditions while overt infections
develop during favourable conditions when bryozoan hosts
are vigorously growing [
]. In addition, we have
experimentally shown that overt infections develop when very
low food levels promote terminal investment , thus
providing further evidence that bryozoans are unable to
suppress infection during adverse conditions.
The adoption of covert infection strategies by
malacosporeans begs the question of whether periods of covert
infection are also employed by other parasites of invertebrate
hosts. Here we discuss this question specifically regarding
their myxozoan relatives - the myxosporeans, whose
lifecycles involve annelids as primary hosts [
]. This focus
enables us to pursue comparative insights and thus to
highlight key questions about covert infection strategies,
impacts and effects. Because covert infections have been
characterised in a range of invertebrate hosts [
], many of
these questions are likely to be equally relevant to covert
infections in other parasite-invertebrate host systems.
We are aware of one study that provides evidence for
a myxosporean undergoing periods of covert infection.
Gilbert & Granath [
] found that actinospores of
Myxobolus cerebralis were released in faeces of
laboratory-infected Tubifex tubifex in a cyclical manner,
approximately 12 times over a 59-day period. This
translated to individual worms releasing actinospores
over several consecutive days punctuated by periods of
no release for a week or more. They also found that
infections could persist for up to 10 months after
actinospore shedding had ceased, that actinospore release could
be resumed some 20 months after worms were first
infected and that individual worms can remain
persistently infected throughout their lifespan. Similar results
for periodic actinospore shedding were obtained for
worms naturally infected in the field. These findings
provide evidence that persistent covert infections of M.
cerebralis may optimise transmission to fish by
periodically converting to overt infections over the multiyear
year life span of worm hosts (e.g. up to 3 years [
The potential for persistent covert infections of M.
cerebralis, however, is apparently not linked with
substantial infection prevalences in the field. PCR-based
assays demonstrate that M. cerebralis prevalences in T.
tubifex populations are usually < 10% and typically < 1%
50, 52, 53
]). Whether such covert infections are
generally avirulent like those characterised for
malacosporeans in bryozoan hosts remains unknown. During
periods of overt infection both malacosporeans and
myxosporeans are virulent, variously causing reduction
in host growth, reproduction (statoblasts in bryozoan
hosts) and mortality [
28, 31, 54–57
Morris & Adams [
] showed that infections of an
unidentified myxosporean can be transmitted when the
oligochaete host, Lumbriculus variegatus, undergoes
fission by architomy (the fragmentation of the body at a
particular point into individual or groups of segments
that then regenerate the missing tissues of the new
]). Similarly, Atkinson & Bartholomew [
report that the myxosporean Myxobilatus gasterostei
transmits during paratomy (the fragmentation of the
body perpendicularly to the antero-posterior axis
following internal tissue reorganisation) of its oligochaete host,
Nais communis. Whether myxosporean infections in
other annelids are transmitted during fission, which has
evolved multiple times in annelids [
unknown. Covert infection strategies evidently do not
depend on the potential for vertical transmission to
products of fission since M. cerebralis undergoes bouts
of covert infection in T. tubifex, a species that does not
undergo fission. Various oligochaetes in freshwater
habitats are known to form cysts that survive adverse
], including T. tubifex [
], but whether
encysted worms harbour covert infections has not been
investigated. It is also unknown if covert infections of
myxosporeans are vertically transmitted to eggs.
Infection of dormant cysts and eggs might enable large-scale
dispersal of myxosporeans, contributing to the dispersal
that is likely to be achieved by ingestion of infected fish
by piscivorous birds [
Baxa et al. [
] provide evidence that some
myxosporean covert infections may result from suppression by
annelid hosts. In their infection trial, clonal lines
deriving from different genetic lineages of T. tubifex were
established in the laboratory and exposed to infection of
M. cerebralis myxospores. Actinospores were released in
varying numbers by clonal lines from different genetic
lineages, but arrested development was revealed by
histological examination and in situ hybridisation in
clonal lines deriving from a particular genetic lineage
characterised by random amplification of polymorphic
DNA (RAPD) PCR analysis. Early stages of development
were present but actinospores were never shed by
worms of this lineage. As far as we know, the extent that
such arrested development may explain differing
susceptibilities to infection associated with genetically distinct
lineages of annelid hosts has not been systematically
addressed. Nor, to our knowledge, is it known whether
infection can be transferred to fish hosts via ingestion of
‘resistant’ worms that harbour covert infections but
suppress actinospore development. Another explanation is
that hyperparasitism may preclude myxosporean
development. Morris & Freeman [
] found that co-infection
by the microsporidian Neoflabelliforma aurantiae results
in hyperparasitism of concurrent myxosporean
infections in oligochaete worms and provide evidence for the
cessation of development and release of actinospores
caused by hyperparasitism. Whether such suppressed
myxosporean infections are capable of subsequent
development and future transmission if microsporidian
infections are lost remains unknown.
The following list of outstanding questions about
the biology of covert infections is inspired by our
specific contrasts between malacosporeans and
myxosporeans. Studies addressing these questions will
increase our understanding of the evolutionary
ecology of parasite-invertebrate host interactions and the
contexts within which covert infection strategies may
evolve and are maintained in hosts with varying life
histories. They are also relevant to the general issue
of how covert infection dynamics may result in
How common are covert infections in invertebrates?
What are the general impacts of covert infections on
Are fission and fragmentation by invertebrate hosts
common routes for covert infection transmission?
Are fission and fragmentation linked with the
evolution of covert infection strategies?
What environmental cues provoke covert infections
to develop into overt infections?
Does cycling between covert and overt infections
characterise covert infection strategies?
Does host capacity to undergo encystment and to
produce resting stages select for covert infection
How common is vertical transmission of covert
infections to sexual offspring?
Do covert infections contribute to parasite dispersal?
How generally do covert infections contribute to the
maintenance and spread of disease reservoirs?
Can covert infections commonly result from
Do covert infections increase the probability of
outcrossing for parasites?
Are covert infections protective to invertebrate
hosts, conferring immunity to subsequent infection
How do covert infection strategies vary in parasites
with simple (direct) and complex (indirect) life-cycles?
How commonly do co-infections suppress
development causing apparent covert infection
and is such developmental suppression
To our knowledge, this is the first long-term study of covert
infections in a field setting and therefore provides unique
insights into what is increasingly being recognised as a
common infection strategy [
]. By addressing a series of
hypotheses, we demonstrate that covert infections are
widespread and persist over space and time in bryozoan
populations. We found variation in prevalences within and
between rivers, at least some of which may be attributed to
variation in environmental conditions (e.g. productivity),
host density and size. The persistence of infected adult
colonies and parasite transmission via statoblasts may
maximise horizontal transmission to fish hosts over time by
establishing a reservoir that poses a long-term risk of
disease outbreaks for farmed fish and wild salmonids. This risk
may be particularly exacerbated by environmental change
]. A review of covert infections caused by myxozoans
in annelid and bryozoan hosts suggests that covert infection
strategies in different systems may share common features.
The implications of covert infections are substantial in
terms of understanding host-parasite interactions and the
impacts of environmental change, and prompt us to
identify several outstanding questions regarding the biology of
Additional file 1: Table S1. Site locations. (DOCX 12 kb)
Additional file 2: Table S2. Environmental variables data. (DOCX 14 kb)
Additional file 3: Detailed CTAB protocol including modifications to the
original protocol. (DOCX 18 kb)
Additional file 4: Figure S1. Water temperature. Temperature
measurements over 12 months according to the 8 sampling trips every
45 days for each river. (TIFF 235 kb)
Additional file 5: Table S3. Water flow data. (DOCX 13 kb)
Additional file 6: Figure S2. PCA plot. Principal components analysis
(PCA) scores for environmental variables. Ellipses are normal contour lines
with probability of 68% done by cluster analysis of rivers. Data points for
each river are coloured by sampling trip (River Avon: 5th trip - 18/04/12; 6th
trip - 11/06/12; 7th trip - 18/07/12; 8th trip - 29/08/12; River Dun: 5th trip
23/04/12; 6th trip - 06/06/12; 7th trip - 23/07/12; 8th trip - 05/09/12).
Variables with vectors pointing in the same direction have similar responses.
Points that are close together correspond to observations that have similar
scores on the components. PC1 explains 56.0% of the variation. PC2
explains 25.4% of the variance. (TIFF 1262 kb)
Additional file 7: Table S4. Summary of PCA analysis on
environmental variables. (DOCX 13 kb)
Additional file 8: Table S5. Colony infection intensity. (DOCX 13 kb)
AIC: Akaike information criterion; BOD: Biochemical oxygen demand;
CFU: Colony forming units; COD: Chemical oxygen demand; DO: Dissolved
oxygen; EA: Environment agency; FTU: Formazin turbidity unit;
GLM: Generalised linear model; GLMM: Generalised linear mixed models;
GV: Granulovirus; HIV: Human immunodeficiency virus; LRT: Likelihood ratio
test; ML: Maximum likelihood estimation; NLS: National laboratory service;
PC: Principal component; PCA: Principal components analysis;
PKD: Proliferative kidney disease; RAPD: Random amplification of
polymorphic DNA; REML: Restricted maximum likelihood estimation;
SE: Standard error; SVD: Singular value decomposition
We thank Nick Taylor (Cefas, UK) and Chris Secombes (University of
Aberdeen, UK) for their contributions to the development and subsequent
advice and support of the NERC Open Case Studentship which funded this
and other related studies. We are grateful to Oliver Robinson (Test Valley
Trout, Ltd., UK) for his generosity and enthusiasm for our work in relation to
PKD outbreaks on fish farms and for his advice about appropriate sites for
our investigations. We are grateful to the following for permission to access
study sites: Darren Butterworth (Trafalgar Fisheries, UK); Robert Starr (river
keeper of the River Dun); Robin Chute (Winchester College, Winchester, UK);
and Mrs. Rochienne Pearce (River Itchen site landowner). We also thank EA
staff for coordinating and conducting water chemistry analyses.
This project was supported by a NERC Open CASE studentship (NE/
019227/1) which received financial contributions from the Centre for
Environment, Fisheries & Aquaculture Science and the Environment
Agency (EA). IF was also funded by SA Pescanova and a PhD research
fellowship from the Fundação para a Ciência e a Tecnologia (SFRH/BD/
86118/2012). None of the funding sources were involved in study
design, data collection, data analysis, data interpretation or in writing
Availability of data and materials
The data supporting the conclusions of this article are included within
the article and its additional files. The datasets generated and/or
analysed during the current study are available in the Dryad repository
BO and HH designed the study. BO, HH and IF conducted field
collections and colony dissections. HH and IF conducted molecular
laboratory work. IF carried out statistical analyses and drafted the
manuscript. CW advised on the nature and coordinated the water
chemistry analyses with the EA. BO, HH and IF actively contributed to
the interpretation of the findings and development of the final
manuscript. All authors read and approved the final manuscript.
Ethics approval and consent to participate
Consent for publication
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in
published maps and institutional affiliations.
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