In situ and low-cost monitoring of particles falling from freshwater animals: from microplastics to parasites

Abstract

A simple and low-cost method of monitoring and collecting particulate matter detaching from (or interacting with) aquatic animals is described using a novel device based on an airlift pump principle applied to floating cages. The efficiency of the technique in particle collection is demonstrated using polyethylene microspheres interacting with a cyprinid fish (Carassius carassius) and a temporarily parasitic stage (glochidia) of an endangered freshwater mussel (Margaritifera margaritifera) dropping from experimentally infested host fish (Salmo trutta). The technique enables the monitoring of temporal dynamics of particle detachment and their continuous collection both in the laboratory and in situ, allowing the experimental animals to be kept under natural water quality regimes and reducing the need for handling and transport. The technique can improve the representativeness of current experimental methods used in the fields of environmental parasitology, animal feeding ecology and microplastic pathway studies in aquatic environments. In particular, it makes it accessible to study the physiological compatibility of glochidia and their hosts, which is an essential but understudied autecological feature in mussel conservation programs worldwide. Field placement of the technique can also aid in outreach programs with pay-offs in the increase of scientific literacy of citizens concerning neglected issues such as the importance of fish hosts for the conservation of freshwater mussels.

Full text

© The Author(s) 2020. Published by Oxford University Press and the Society for Experimental Biology.
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/
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Volume 8 • 2020 10.1093/conphys/coaa088
Toolbox
In situ and low-cost monitoring of particles falling
from freshwater animals: from microplastics to
parasites
Karel Douda1,*, Felipe Escobar-Calderón1,BarboraVodáková
1,PavelHorký
1,Ond
ˇ
rej Slavík1
and Ronaldo Sousa2
1Department of Zoology and Fisheries, Czech University of Life Sciences Prague, Kamýcká 129, CZ-165 00, Prague, Czech Republic
2CBMA, Centre of Molecular and Environmental Biology, Department of Biology, University of Minho, Campus Gualtar, 4710-057 Braga, Portugal
*Corresponding author: Department of Zoology and Fisheries, Czech University of Life Sciences Prague, Kamýcká 129, Prague CZ-16500,
Czech Republic. Email: k.douda@gmail.com
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A simple and low-cost method of monitoring and collecting particulate matter detaching from (or interacting with) aquatic
animals is described using a novel device based on an airlift pump principle applied to oating cages. The eciency of the
technique in particle collection is demonstrated using polyethylene microspheres interacting with a cyprinid sh (Carassius
carassius) and a temporarily parasitic stage (glochidia) of an endangered freshwater mussel (Margaritifera margaritifera)
dropping from experimentally infested host sh (Salmo trutta). The technique enables the monitoring of temporal dynamics
of particle detachment and their continuous collection both in the laboratory and in situ, allowing the experimental animals to
be kept under natural water quality regimes and reducing the need for handling and transport. The technique can improve the
representativeness of current experimental methods used in the elds of environmental parasitology, animal feeding ecology
and microplastic pathway studies in aquatic environments. In particular, it makes it accessible to study the physiological
compatibility of glochidia and their hosts, which is an essential but understudied autecological feature in mussel conservation
programs worldwide. Field placement of the technique can also aid in outreach programs with pay-os in the increase of
scientic literacy of citizens concerning neglected issues such as the importance of sh hosts for the conservation of freshwater
mussels.
Key words: Aquatic animals, drop-o, sh, freshwater mussels, glochidia, host–parasite relationships, microparticles, microplastics
Editor: John Mandelman
Received 22 April 2020; Revised 17 June 2020; Editorial Decision 9 September 2020; Accepted 9 September 2020
Cite as: Douda K, Escobar-Calderón F,Vodáková B, Horký P, Slavík O, Sousa R (2020) Insitu andlow-cost monitoring of par ticles falling fromfreshwater
animals: from microplastics to parasites. Conserv Physiol 8(1): coaa088; doi:10.1093/conphys/coaa088.
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Introduction
The inherent complexity of various ecological processes war-
rants the efficient combination of laboratory and field exper-
iments. However, despite the rapid development of tools
designed to enhance field data collection (e.g. remote elec-
tronic control systems; Burnett et al., 2013;Wilson et al.,
2014;Kubizˇnák et al., 2019) there still exist many research
areas where no field-based, low-cost technical solutions are
available for primary data collection. This situation is espe-
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Too l bo x Conservation Physiology • Volume 8 2020
cially true for aquatic organisms, which may restrict the
collection of data for many ecological and physiological pro-
cesses to laboratories or during short-term invasive sampling
campaigns (e.g. Barber et al., 2008;Hart et al., 2018).
Diverse research fields such as ecological parasitology,
ecotoxicology, aquatic animal nutrition and reproductive
biology require techniques to collect objects detaching from
live aquatic animals. Laboratory methods exist for collecting
parasite stages (Dodd et al., 2005;Marchiori et al., 2013),
faecal pellets (Shomorin et al., 2019) or eggs (Gonsar et al.,
2012) in recirculating systems on screens. These methods
make the following possible: the study of the time course
of particle detachment at the individual level, the evaluation
of the daily feed intake of animals in aquaculture facilities
and the collection of particles over extended periods of time.
The collection of fallen particles has also proven essential for
understanding various aspects of aquatic animal physiology
such as digestibility analyses (Da Mota et al., 2015;
Dvergedal et al., 2019) and host–parasite compatibility (e.g.
Rogers-Lowery et al., 2007;Dodd et al., 2005;Donrovich
et al., 2017). However, laboratory approaches have the
disadvantage of being limited to a range of model organisms
for which long-term holding under artificial conditions has
been mastered (Levy and Currie, 2015;Russell et al., 2017).
Consequently, a lack of data persists for most animal species
in which the laboratory approach is not feasible because it can
inadvertently affect their behaviour, biological rhythms and
physiology (Calisi and Bentley, 2009). This, coupled with the
high operational costs and labor requirements of the research
facilities needed, makes the laboratory approach unsuitable
in many areas of ecology and conservation physiology
research.
Here, and as a proof of concept, we describe a new tech-
nique that can be used in the field to collect objects detaching
from (or interacting with) aquatic organisms using a flow-
through cage system. For this, we assessed the technique’s
efficiency to collect (i) microplastics (polyethylene micro-
spheres) and (ii) juveniles (post-parasitic stage) of an endan-
gered species. We choose these two cases because in one hand
microplastics have gain traction as a recent relevant research
topic due to the possible deleterious effects on consumption,
growth, reproduction and survival of aquatic animals (Foley
et al., 2018). However, the level of knowledge in freshwater
ecosystems lags behind what has been explored in marine
ecosystems (Eerkes-Medrano et al., 2015), and the interaction
between microplastics and freshwater organisms is particu-
larly understudied for wildlife compared to laboratory models
(de Sá et al., 2018). On the other hand, and given their
complex life cycle, we used one species of freshwater mus-
sels (Bivalvia: Unionida), one of the most threatened faunal
groups in the planet, which in the past decades has been highly
studied and subjected to several conservation management
plans including captive breeding programs (Lopes-Lima et al.,
2017;Ferreira-Rodríguez et al., 2019). This group of bivalves
has a temporarily parasitic larval stage (glochidium; size,
50–400 μm) that must attach to the body surface of a suit-
able fish and become encapsulated in the epithelial layer to
metamorphose into a juvenile mussel (Kat, 1984;Modesto
et al., 2018), then it ruptures the capsule and detaches from
the host. Freshwater mussel–fish relationships have become
useful models for addressing questions in fish ecology (Gopko
et al., 2018;Horký et al., 2019;Methling et al., 2019),
toxicology (Defo et al., 2019;Douda et al., 2019) and the
conservation biology of host–affiliate relationships (Tremblay
et al., 2016;Schneider et al., 2017).
Various laboratory methods have been established for
the study of the metamorphosis success rate of glochidia
using adapted multi-unit laboratory fish-holding recirculation
systems (Dodd et al., 2005;Hazelton et al., 2013;Douda
et al., 2018;Dudding et al., 2019), sets of aquaria adapted for
periodical or continuous siphoning (Reis et al., 2014;Douda
et al., 2014;Reichard et al., 2015;Donrovich et al., 2017)
or other custom-made fish holding tanks (Taeubert et al.,
2013;Eybe et al., 2015;Huber and Geist, 2017;Soler et al.,
2018). However, some of these methods can be problematic
(especially when used for fish collected in the field), leading
often to high fish mortality during experiments (Taeubert
et al., 2013;Huber and Geist, 2017;Soler et al., 2018), reduc-
ing the representativeness of the results. The fact that there
is currently no available method for the collection of mussel
juveniles falling from the fish host under field conditions
strongly limits our ability to test new potential hosts in species
where transport to the laboratory is problematic, or in areas
without suitable laboratory infrastructure. Such limitation is
one of the main reasons for the insufficient knowledge of the
host sources of freshwater mussels (Modesto et al., 2018) and
for the need to look for new methods that are feasible without
a laboratory (Hart et al., 2018).
Given the above-described background and the need to
develop simple methods that increase information about
basic autecological processes, the main aim of this study was
to describe a low-cost technique that may be employed in
several ecological topics related to conservation physiology
of aquatic animals (from simple assessment of animal-
microplastics interactions to more complicated analysis of
host–parasite relationships). We also discussed the use of this
technique in other topics, including outreach programs.
Methods
To demonstrate the utility of the technique in real-world
ecological problems, we present two examples that can be
performed with this device, whether it is in a laboratory or
a field. The first quantifies the interaction time and capture
efficiency of the device for externally added standard par-
ticles in the laboratory with potential use in the study of
animal–microplastics interactions. The second illustrates a
breakthrough advance in field-based fish–glochidia interac-
tion studies by addressing questions previously tractable only
under laboratory conditions.
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Conservation Physiology • Volume 8 2020 Too l bo x
Principle and construction of the device
Floating board and cages
The floating drop-off particle collector (FDPC) unit operates
on a free-floating board (width: 50 mm; polystyrene) weighed
down from the upper and bottom side by protective sheets
(thickness: 5–10 mm; polypropylene). Five animal holding
tanks are suspended below the floating board, each posi-
tioned within five different divisions (Figs 1 and 2). The
divisions are created by heat welding 5–10 mm polypropylene
sheets perpendicular to the main floating board at regular
intervals. The bottom of the tank lies on a single sheet (thick-
ness: 5 mm) to which the perpendicular sheets of the divisions
are heat welded. In each division, an experimental tank is
placed to form a cage for the fish. Commercially available
boxes with a smooth and undiversified internal surface can
be used. Here, polypropylene fish tanks (volume: 20 L, length
x width x height: 34 x 22 x 28 cm; T-Box S, Keter Italia S.p.A.,
Italy) were used. To firmly fit the tanks into each division and
allow passage of water from the exterior, a gap between the
floating board and the tanks in each division was created by
inserting two silicone blocks (height: 12 mm) with smooth
edges to prevent injury to the fish. The dimensions of the
silicone blocks need to be adjusted to the size of the organisms
tested.
Air and water ow
The FDPC device operates using the principle of airlift pump-
ing. Each tank is equipped with its own riser pipe (diameter:
20 mm; PVC pipe), the pressured air required by the units
during operation is provided by land-positioned compressors.
The air is injected into the bottom part of the riser pipes in
each holding tank, and because the mixture of air and water
is less dense than the surrounding water, it rises to the top
aperture, sucking water and solids from the bottom of the
tank and transporting them to the collection net positioned
above the main floating board. The riser pipe outlet in the top
of the FDPC is connected to a 90-degree bend, ending 130 mm
above the water surface level (80 mm above the floating board
surface), just above a collecting filter cylinder. The main air
supply line starts with an electrical air compressor to which a
hose (inner diameter: 135 mm) is connected. The other end of
the hose is attached to one end of the FDPC device on top of
the floating board. From there, a manifold air divider valve
distributes the air to the different riser pipes (or is left open
to stabilize the airflow if needed—see below) through 4 mm
(inner diameter) silicone tubes. Each tube is equipped with a
two-way air control valve. A single air compressor can feed
several FDPC units; here, one 100-W compressor (airflow:
110 L min−1; air pressure: 0.035 MPa, 102 W; Hailea ACO-
009, China) was successfully used to feed 2–3 FDPC units.
Because the flow rate determines the entrapping effectivity
of the pump, it is necessary to measure the water flow through
each filter and standardize it among tanks. The water flow
through the outlets can be measured by a graduated collection
Figure 1: Side (A), front (B) and top (C) schematic view of FDPC. r,
riser pipe; s, lter cylinder; f, feeding and calibration port; b, oating
board; c, polypropylene cage; a, air delivery hose; red arrows, water
ow; blue arrow, air ow.
vessel placed where the filter cylinders are usually located and
adjusted by changing the amount of air being pumped to the
riser pipes using the two-way air valves connected to the air
tubes for each tank. The mean ±SD water flow through the
individual tanks under the above-described settings during
both experiments was 45.6 ±9.6 mL s−1.
Collecting cylinders
The collecting filter cylinders (Fig. 2C) are made from PVC
pipe (diameter: 115 mm; height: 65 mm) with a nylon screen
of specific mesh adjusted to the size of the monitored particles
attached (here, we used a loop size of 139 μm). The filter
cylinders are placed into PVC positioning box fixed on top
of the FDPC, which stabilizes the position of the filter at
the desired angle against the riser pipe outlet (we used 45
degrees as the optimal angle). The height of the openings in
the positioning box determines the water level around the
cylinder and allows the presence of a pool of water above the
bottom part of the screen. This pool keeps the particles under
water after recovery if needed. Alternatively, other type of
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Too l bo x Conservation Physiology • Volume 8 2020
Figure 2: Example use of the FDPCs for the sampling of Margaritifera
margaritifera juveniles dropping from host sh (Salmo trutta): (A)
polypropylene structure arrangement for a 5-cage system, (B) eld
deployment of 7 systems with 34 x 22 x 28 cm cages and (C) detail of
the collection cylinder.
screens, such as wedge wire screens, can be used, if necessary,
to keep the recovered material out of the water (not tested
here, see Shomorin et al., 2019 for details).
Feeding and calibration port
The FDPC is equipped with a set of additional ports located
in each section opposite to the main riser pipe. A silicone
tube (inner diameter: 10 mm) is positioned in each opening
(ending 5 mm above the floating board). The function of
these apertures (hereinafter feeding and calibration ports) is
to allow the introduction of external food items during the
experiment (if needed) or a known number of particles of
interest for exposure or calibration purposes.
The cost of the system described for all experiments was
approximately $1105 per 35 tanks distributed among 7 FDPC
units (see Table S1 for a detailed description). The system can
be easily built using an electric saw, plastic welding heat gun
(with compatible polypropylene rods), electric screwdriver
and drill bit and moved to any water body with available
electricity on the bank. While we used plug-in compressors
and a 230-volt power connection, solar or battery sources
alongside voltage converters can be used to make the system
more portable. The device does not require any construction
of solid structures or racks and adapts to possible fluctuations
in water level.
Proof of concept
Example 1: polyethylene microspheres
Cyprinid fish Carassius carassius (Linnaeus, 1758) individ-
uals (mean total length: 127 mm; mean body mass: 34 g)
obtained from a laboratory breeding population at the Czech
University of Life Sciences Prague (Czech Republic) were kept
in a 250-L aquarium at 15◦C, and a light–dark regime of
12:12 h before the start of the experiment. Fish were fed daily
with commercial fish pellets (Pond Pellet, 5–6 mm; Tetra,
Germany) before and during the experiment. A FDPC unit
was installed in a 200 x 100 x 100 cm (length x width x
height) laboratory tank with dechlorinated tap water (1800 L)
under identical temperature and photoperiod conditions as
described above. On the day of the start of the experiment,
five randomly selected fish were extracted from the aquarium
and placed into each of the tanks of the FDPC.
The microplastics, Red Polyethylene Microspheres
(1.12 g cc−1, 500–600 μm), were purchased from Cospheric
(Santa Barbara, CA, USA). To prevent the particles from
floating or creating clumps, an organic food-grade surfactant
(Tween 80 Biocompatible Surfactant, Cospheric, CA, USA)
was used. The microplastics (106–114 particles) were
introduced into the respective tanks in the FDPC with the
help of a syringe attached to a silicone tube. The assessment
of flushed particles was performed at 1, 6, 24, 48, 72, 96,
120, 144 and 168 h after the start of the experiment. At
each time, the five collecting cylinders of the FDPC were
replaced with new clean filters, and the used filters were then
observed under a microscope to assess the number and status
of particles recovered.
Example 2: parasitism success in an
endangered species
A second experiment applied the FDPCs to monitor para-
sitism success and to collect juveniles of the freshwater mussel
Margaritifera margaritifera (Linnaeus, 1758) detaching from
its fish host, Salmo trutta Linnaeus, 1758. It should be noted
that previous field studies have been restricted to evalua-
tion of glochidia attachment intensity observed on wild fish
(Salonen et al., 2017;Dias et al., 2020), whereas evaluation
of metamorphosis success has been limited to laboratory
studies (e.g. Douda et al., 2017;Schneider et al., 2017). The
success of M. margaritifera parasitization was tested using
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Conservation Physiology • Volume 8 2020 Too l bo x
larvae from two different source populations (with different
qualities of glochidia) experimentally infesting host fish from
two different populations.
For this, the experimental S. trutta were caught by elec-
trofishing (650 V, 4 A, pulsed D.C.) in two streams (popu-
lation Fish-A, ˇ
Zivný potok stream, 49◦239N, 14◦132E;
population Fish-B, ˇ
Castá stream, 48◦554N, 13◦4027 E)
within the Vltava River basin (Czech Republic) with no
current M. margaritifera populations. The fish were anaes-
thetized with 2-phenoxy-ethanol (0.2 mL L−1; Merck KGaA,
Germany), measured (total length: mean 164 mm, range
95–216 mm), weighed (body mass: mean 40 g, range 6–90 g)
and individually marked 6–13 days before the infestations.
Passive integrated transponders (PITs; Trovan ID100, 0.1 g in
air, 12 ×2.1 mm; EID Aalten B.V., Aalten, the Netherlands)
were inserted into the dorsal muscle using a syringe. After
marking, the fish were kept in side-arm of the Vltavský potok
stream (48◦590.5”N, 13◦3938E) in the ˇ
Sumava National
Park hatchery before infestation with glochidia.
Parasitic glochidia of M. margaritifera were obtained from
female mussels sampled from two different populations in the
Vltava River basin (Czech Republic) (population Gloch-A,
Blanice River, 48◦5534N, 13◦5812E; population Gloch-
B, Malˇse River, 48◦3901.5N14
◦2800.3 E). To obtain
glochidia of M. margaritifera, several mussel individuals
were monitored in the field and when glochidia release was
observed, the individuals were collected and placed into a
shallow 5-L vessel to stimulate further glochidia release. The
clumps of glochidia released were extracted with the help of
a pipette and observed under a microscope to assess viability.
Then, the glochidia were transferred to 5-L containers with
river water. Two separate mixtures of glochidia obtained
from 35 and 3 female mussels from populations Gloch-A
and Gloch-B, respectively, was used (August 2018, 7 days
between the two infestation events). After the glochidia
were extracted, the females were returned to the same
collection location. The containers with glochidia were
transported immediately in cooling boxes to the Vltavský
potok stream, where the infestations were performed in the
same day.
Fish were infested with glochidia in August 2018 in a
common bath suspension with densities of 15 400 ±3666 and
11 200 ±3516 (mean ±SD) glochidia L−1for populations
Gloch-A and Gloch-B, respectively. Density was assessed
by counting ten 1-mL subsamples. The viability of the
glochidia was tested by evaluating their snapping action in a
NaCl solution immediately before infestation (Roberts and
Barnhart, 1999). The average percentage of viable (reacting)
glochidia in the inoculation bath was 31% in Gloch-A and
74% in Gloch-B. The infestation procedure lasted 15 min,
and the density of fish in the glochidia suspension was
1 fish L−1. Individuals from both fish populations were
infested in a common bath. The control (uninfested) fish
were treated with the same handling procedures (i.e. transfer
between baths). After infection, the fish were released into
a seminatural side-arm of the Vltavský potok stream with
a natural gravel/sand bottom (length: 47 m; width: 2–3 m;
depth: 0.1–0.6 m) and an adjacent earth pond (area: 139 m2;
max. depth: 1.5 m).
The monitoring of falling juvenile mussels using FDPCs
was initiated upon reaching the sum of temperatures reported
as usual for the start of juvenile mussels dropping from host
fish (Hruˇska, 1992), which occurred in June 2019 (total
number of days from infestation: Gloch-A, 310 and Gloch-
B, 317). The average daily temperature during the whole
period ranged between 0.2 and 16.1◦C, and the total sum of
daily degrees until placement in the FDPCs ranged between
1573 and 1783. Seven FDPC units (total: 35 holding tanks)
were placed directly at the site where the fish had spent the
previous part of the parasitic period. The fish were caught
as described above and were gradually placed in the FDPCs,
where they spent 6–8 days at average daily temperature
during monitoring 13.2 ±1.0◦C (range: 12.0–15.1◦C). The
relative body weights (condition factors) of 12 randomly
selected fish individuals were determined using the equation
K = 100 x somatic weight (g)/(standard length [cm])3before
the placement and after the removal of the FDPCs. We
have verified the functionality of the feeding and calibration
ports for live feeds but did not add food items on a regu-
lar basis because the presence of live aquatic invertebrates
(mayfly larvae, benthic crustaceans) was regularly detected
on the filters, indicating natural food being supplied to the
tanks in this experiment. The FDPC collecting cylinders were
exchanged at 1–2-day intervals and inspected at 10–40x
magnification under the microscope. Juvenile mussels falling
from the hosts were classified as live if valve or foot movement
was observed. The average rate of parasite detachment from
fish (number of juvenile mussels day−1g−1of fish body
weight) was determined together with the success rate of
metamorphosis during the monitored period (the percent-
age of dead and live juveniles falling from the fish). Fish
individuals were returned to their site of capture after the
experiment.
To verify the temperature conditions in the FDPCs, a
datalogger (temperature accuracy: 0.1◦C; Hobo, Onset, USA)
was placed inside and outside the device, recording data every
15 min for 7 days. For the field flushing efficiency test,
uninfected control fish were placed in 3 tanks of an FDPC,
and 36–74 mussel juveniles were then placed inside the unit
using the feeding port. For the next 96 hours, monitoring
was performed as described above to determine the success
of recapture.
We used paired Wilcoxon rank-sum tests to determine
whether the detachment rate of juveniles and metamorphosis
success (arcsine-transformed proportion of viable juveniles)
differed between the different host–parasite population com-
binations. Paired t-tests were used to compare fish condition
factor and temperature differences. All analyses were per-
formed in R 3.5.2 (R Core Team, 2019).
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Results and discussion
Example 1: polyethylene microspheres
Mortality of C. carassius during the experiment was zero
and there were no signs of skin or fin injuries. The capture
efficiency using polyethylene microspheres showed a mean
(±SD) particle flushing efficiency of 95.9 ±4.5%. Most of
the particles (91.3 ±5.1%) were f lushed in the first 24 h, and
the last particles were recovered 120–144 h after insertion
(see Table S2 for details). Microspheres recovered in the later
stages of the experiment (72–144 h) showed that they had
been mechanically damaged and small particle fragments
were also present. Although it was not specifically studied
here, the relatively long residence time and physical damage
of these particles indicated that they had passed through the
digestive tract of the fish and were harmed by the pharyngeal
teeth.
By capturing particles leaving the enclosure space, the
device allows determining the time and concentration of
exposure to particles while being held under ambient envi-
ronmental conditions. The availability of well-defined (colour,
size, relative density, shape) plastic particles for experimental
purposes enables this to be done effectively and offers new
experimental possibilities. In addition, water flow through the
system can be regulated to adjust the residence time.
Example 2: parasitism success in an
endangered species
Host fish (S. trutta) mortality was zero, there were no signs
of skin or fin injuries, and the condition factor of fish did not
change (P>0.05) during the experiment. The flushing effi-
ciency of the M. margaritifera juveniles in the field (recapture
rate of added juveniles) ranged between 88.1% and 100.0%,
and 90.4% to 98.6% of juveniles were recovered within the
first 24 hours. There was no difference in the temperature
recorded inside and outside the devices (P>0.05; mean dif-
ference ±SD: 0.05 ±0.08◦C).
The estimated average M. margaritifera juvenile detach-
ment rate across all fish was 0.16 ±0.47 juveniles day−1g−1,
and the average percentage of successfully metamorphosed
glochidia was 74.0 ±30.2%. In terms of the detachment rate,
there were significant differences between the fish infested
with different mussel populations (Fig. 3A). Fishes infested
with Gloch-A had a significantly (P<0.001) lower juvenile
detachment rate (0.01 ±0.02 juveniles day−1g−1) than fish
infested with Gloch-B (0.68 ±0.79 juveniles day−1g−1); but
there were no detectable differences in the juvenile detach-
ment rate between host fish populations (P>0.05).
In terms of juvenile mussel metamorphosis success, a
slightly higher percentage of live juveniles was associated
with the fish infested with Gloch-B (78.6%, versus 70.4%
for Gloch-A, Fig. 3B), which corresponds with the higher
detachment rate in this fish population, but no significant
Figure 3: (A) The rate of Margaritifera margaritifera juvenile
detachment per gram of sh body weight (pairwise Wilcoxon test,
dierences between mussel populations—P<0.001, n=4–18) and
(B) the proportion of successfully metamorphosed glochidia during
the 14-day monitoring period (1573–1783 degree days from
infestation; pairwise Wilcoxon test, all P>0.5, n=4–18) as detected
by the FDPC. The median, interquartile range and min/max for
dierent combinations of source populations of parasites (red/blue)
and hosts (hatched/unhatched) are displayed.
differences were detected (all P>0.05). A total of 2377
detached M. margaritifera juveniles were sampled.
These results show that FDPC is able to detect differences
in the physiological compatibility of different combinations
of source glochidia and host populations. In our case, the
results demonstrate a greater efficiency in the use of S. trutta
hosts by the glochidia from population B possibly due to
immunological mechanisms (Rogers-Lowery et al., 2007), or
due to a lower quality of glochidia produced by population
A (indicated also by the initial viability analysis, see above), a
common problem in freshwater mussel propagation activities
(Patterson et al., 2018). In terms of conservation application,
it shows us which mussel population provides a more efficient
source of glochidia for possible rescue or bioindication breed-
ing. On the other hand, the results do not indicate a different
ability of the two fish strains to host M. margaritifera due to
local adaptation as recorded by previous studies (e.g. Douda
et al., 2017;Schneider et al., 2017). Although a more complex
study design would be needed to take into account the effect
of glochidia viability, and test the effects of recorded lower
metamorphosis success rates in the combination of Fish-B
and Gloch-A populations, both populations can be considered
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Conservation Physiology • Volume 8 2020 Tool box
physiologically compatible hosts. Therefore, the FDPC allows
addressing the geometry of local adaptations between mussels
and fish by studying metamorphosis success directly in the
field (in remote geographical locations, under natural temper-
ature and photoperiod regimes and water quality conditions),
which to our knowledge has not been possible before.
General discussion and way forward
This study described the construction of a field-deployable
floating device for the continuous monitoring of detachment
or interaction regimes of particles associated with aquatic
animals. This novel approach is cheap and mobile, and can
be used in other type of environmental studies (e.g. faeces-
based molecular diet analyses and ingested microplastic quan-
tification) (Nelms et al., 2019) using fishes and other aquatic
animals (e.g. crayfish and other macroinvertebrates, amphib-
ians).
The use of nonlethal methods to collect fish faeces from
animals exposed to microplastics can prove to be a valuable
addition to this type of study (Hoang and Felix-Kim, 2019;
Kazour et al., 2018), allowing us to record the dynamics of
microplastic excretion. The device can be especially useful
when a long transport distance would be necessary and
risky, or when the acclimatization to the available laboratory
conditions is problematic (Calisi and Bentley, 2009). It should
be highlighted that this method cannot be easily used for non-
specific monitoring of plastics in the field due to their possible
source from the surrounding environment and the device itself
(Löder and Gerdts, 2015;Li et al., 2018). On the contrary,
the proposed use tested here as proof of concept consists on
the controlled exposure of organisms to plastics of specific
properties and detectability (Shim et al., 2017;Heinrich et al.,
2020) either before or during (as showed here) the placement
into the system and monitoring the regime of particles-animal
interaction under natural water quality and temperature and
photoperiod regimes.
Although we showed that the FDPC is ideal for collecting
particles dropping from or interacting with fish in laboratory
and oligotrophic habitats, slight alterations to the presented
system can further increase its range of applications. The use
of wedge screens can be suitable for the collection of faeces
more effectively (Dvergedal et al., 2019;Shomorin et al.,
2019), and the system can be surrounded by protective nets
to prevent input of other prey items (when providing food
items manually) or other types of potential interference (e.g.
in the case of filter-feeders). Another possibility to extend the
usability of the system is the implementation of technolog-
ical accessories to record and report online the behavioural
activity of the objects studied, environmental conditions and
system malfunctions, which has not been possible without
continuous operator presence until recently (Kubizˇnák et al.,
2019;Sheehan et al., 2020).
Another promising opportunity for the FDPC application
is the conservation biology of freshwater mussels, which are
declining worldwide (Lopes-Lima et al., 2014,2018). The
use of FDPCs in mussel conservation can involve two main
activities. First, as demonstrated here using M. margaritifera,
the FDPC represents a cheap, reliable, and deployable mean
of testing the glochidia metamorphosis success rate—a critical
knowledge for the determination of conservation units and
host resource management (Modesto et al., 2018). Second, the
FDPC can be a powerful tool for the recovery of both larvae
and juveniles from endangered freshwater mussels. The use of
this (or similar) techniques to increase our knowledge about
basic autecological features of freshwater mussels is highly
welcome, because it has been shown that adult mussels held
in the laboratory conditions over long terms exhibit lower
growth, altered metabolism and higher mortality (Patterson
et al., 2018;Roznere et al., 2014).Although a great increase in
the number of studies addressing ecological and conservation
issues of freshwater mussels can be found in the past decades,
the reality is that basic information on key autecological (e.g.
distribution, density, population size structures) features are
still lacking (Lopes-Lima et al., 2020) especially in some areas
where equipped laboratories or personal are not available.
In fact, one key information gap is their reproduction and
the metamorphosis of glochidia to juveniles. The device and
methodology described here can overcome some of the bias
(water quality, feeding and temperature differences) already
described in the usual laboratory procedures and can help to
expand this type of research into new geographical areas.
The device also has good potential for use in other biotic
interactions. For example, Trematoda parasites produce in
their intermediate (molluscan) host free-living larvae (cer-
cariae), which swim actively or float passively in the water to
find and infect the next host. An important branch of aquatic
parasitology is the estimation of cercarial production. This is
challenging in field conditions, because so far, the only way to
estimate cercariae production has been to place the mollusc in
a container for a period of time to be able to count the larvae
(e.g. Taskinen 1998). The FDPC system described here can be
an important innovation in this type of research. In addition,
the possibility of placing the system in a freely accessible
(compared to a remote and quarantined laboratory) location
in the field can be beneficial for educational purposes. In the
case of our field site near the fish hatchery of ˇ
Sumava National
Park, there were many opportunities to demonstrate the
device to students and other visitors and thus communicate
the fish-mussel host-parasitic system and their importance for
conservation research programs.
Despite the possible advantages, it is important to take
into consideration that although the device can be located
in a river or a lake, it is not a physically natural habitat but
an enclosure. Thus, it brings an effective advantage in some
fundamental parameters (temperature and light regimes, and
water quality), but on the contrary, it does not allow a
number of natural behaviours (e.g. movements of animals to
foraging areas or an interaction with substrate). Therefore,
in particular cases, it will be necessary to determine whether
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Too l bo x Conservation Physiology • Volume 8 2020
the caging can affect the studied parameter. In the same vein,
and although the device can eliminate the need of organisms
transport over long distances and reduce the risks of disease
transfer to or from laboratories or among catchments, as a
field-deployable device, the FDPC itself could contribute to
the movement of diseases and species. Because of this, we
strictly recommend that all parts of the FDPC in contact
with water must be disinfected and allowed to dry completely
before being transported to another location.
In conclusion, collecting particles dropping from aquatic
animals directly in the field not only provides opportunities
to greatly increase the volume and type of data that can
be collected in environmental parasitology or animal feeding
ecology, but also enables the acquisition of new types of data
in emerging research fields, such as microplastic pathway
studies. Further research is needed to test FDPCs in other
water systems and in association with other research topics.
This system has excellent prerequisites for interconnection
with remote electronic monitoring systems. Continued tech-
nological advances will make field-deployed floating systems
an increasingly viable and versatile option without needing
a sophisticated laboratory for holding organisms originat-
ing in the wild with the associated long-distance transport.
The simple and low-cost design, field accessibility and easy
operation also allow its use in outreach programs, increasing
the scientific literacy of citizens in very specific topics such
as the importance of fish to conserve critically endangered
freshwater mussels.
Supplementary material
Supplementary material is available at Conservation Physiol-
ogy online.
Author contributions
K.D. conceived the idea and designed the hardware. F.E-C.,
B.V. and K.D. performed the calibration, laboratory and field
experiments. P.H., O. S. and K.D. collected the fish hosts
and deployed the field units. All authors provided critical
feedback, participated in manuscript writing and approved
the final manuscript.
Funding
This work was supported by the Czech Science Foundation
[19-05510S] and the European Regional Development Fund
[CZ.02.1.01/0.0/0.0/16_019/0000845, CZ.05.4.27/0.0/0.0
/15_009/0004620].
Acknowledgements
We t ha nk Zb y n ˇek Janˇci and Bohumil Dort for the help in
the field, the nature conservation authorities for providing
permits and access to the research area in Borová Lada and
two anonymous reviewers for their helpful comments on an
earlier draft. All experiments were in compliance with the
current laws of the Czech Republic Act No. 246/1992 coll.
on the protection of animals against cruelty.
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