Introduction
Plastic is one of the most abundant synthetic substances on the planet
(Lambert et al., 2014). An estimated 381 million tons of plastics were
produced in 2015 and this number keeps rising (Ritchie & Roser, 2018).
Widely recognized since the 1950s as cheaper, lighter and more durable
than other materials such as metal and glass, plastics (most commonly
polystyrene (PS), poly(ethylene terephthalate) (PET), polyurethane
(PUR), poly(vinyl chloride) PVC, polyethylene (PE) and polypropylene
(PP); (Strungaru et al., 2019)) have become the primary material in
disposable packaging, and are pivotal in the construction and automotive
industries; applications are practically endless (Lambert et al., 2014).
However, plastics accumulate in the environment (Chamas et al., 2020)
and are emerging as a matter of concern in both aquatic and terrestrial
systems (Andrady, 2011; de Souza Machado, Kloas, et al., 2018; Rillig,
2012).
Microplastics (MPs) are defined as any plastic particle <5 mm
in diameter (Hidalgo-Ruz et al., 2012). They are produced either
intentionally (e.g., by the cosmetic industry for use as skin
exfoliants) or form through the breakdown of larger pieces (Wang et al.,
2019). Weathering of plastic particles in the environment can occur via
microbial action (Yuan et al., 2020), photodegradation by UV rays
(McKeen, 2013), chemical action (Liu et al., 2020) and heat (Andrady,
2011; Pischedda et al., 2019; Wang et al., 2019). Microplastics can
cause both physical and biotic effects in soil. Physical effects include
changes in soil bulk density and structure, which can alter water
holding capacity and nutrient availability (de Souza Machado, Lau, et
al., 2018). In freshwater systems, MP particles can undergo changes in
hydrophobicity and buoyancy, which can make them more bioavailable to
certain organisms (Helcoski et al., 2020). Concentrations of MPs in soil
have been observed to vary widely, ranging from < 0.01
items/kg to > 10,000 items/kg (Jacques & Prosser, 2021)
and they can be as high as 7% by volume in some of the most polluted
top-soils where plastic mulches and biosolids are used (Fuller &
Gautam, 2016). However, pinpointing environmentally relevant
contamination levels remains difficult at this time, as publicly
available large-scale terrestrial monitoring data and standardized
quantification campaigns are limited or lacking altogether. In addition,
a focus should be on potentially future, higher levels of this
contaminant, at least within a global change biology framework (Rillig
et al., 2021).
Although much of the study of environmental impacts of MPs in
terrestrial systems has been focused on macrofauna (Prokić et al.,
2021), such as birds (Azzarello & Van Vleet, 1987) and mammals (Zantis
et al., 2021), new evidence shows that smaller soil organisms are also
affected (Ren et al., 2021; Wang et al., 2019). For example, as much as
73% of springtails can ingest MPs < 2 μm, which contributes
to slowing down their movement (Kim & An, 2020). In another study,
three nematode species (Caenorhabditis elegans ,Acrobeloides nanus and Plectus acuminatus ) could be seen
ingesting MPs < 1 μm (Mueller et al., 2020), and the structure
of soil microbial communities can also be affected by MPs (Huang et al.,
2019; Zhang et al., 2019).
Protists are a diverse group of primarily microscopic, unicellular
organisms that are abundant in both aquatic and terrestrial ecosystems
(Adl et al., 2005; Geisen et al., 2018). A single gram of dry soil can
contain 104 - 107 active individuals
spanning from photosynthetic (algae), heterotrophic (phagotrophic), and
mixotrophic (i.e., those capable of both photosynthesis and phagotrophy)
protists. As such, protists are a pivotal, often neglected, component of
the soil microbiota playing important roles in carbon and nutrient
cycling (Adl & Gupta, 2006, Geisen et al., 2020). Phagotrophic protists
ingest a variety of food sources including bacteria, fungi, plants and a
range of mesofauna and other protists (Geisen et al., 2018). Many are
free living and actively utilize locomotion by way of cilia or flagella
to locate and capture prey and other resources, which they ingest
through their oral groove into food vacuoles (Verni & Gualtieri, 1997).
Protists can serve as bioindicators of soil contamination. They have
been used in toxicological studies as model species, for example to
assess metal toxicity of cadmium and zinc (Johansen et al., 2018). Since
all phagotrophic soil protist species are transparent, they are
excellent candidates for use as indicator species as putative toxic
particles can easily be seen internally. Despite this potential and
their abundance in soils, the ecology of protists and their role in soil
microbial communities is still poorly understood (Rillig & Bonkowski,
2018).
Whether soil phagotrophic protists in general can ingest MPs is unknown. No papers exist
that highlight both MP and soil protists specifically. A search for
“microplastic*” on ‘Web of Science’ on December 3rd,
2020 resulted in 5531 papers. Searching for “Protist*” within
“microplastic*” returned only three papers, two of which were focusing primarily on aquatic systems and the other one was a call for research on soil
protists and MPs (Rillig & Bonkowski, 2018). The fact that they are an
important energy channel in the soil food web means that protists can
serve as vectors of MPs to higher trophic levels (Setälä et al., 2014).
This hypothesis is plausible considering that other soil microorganisms
with equivalent mechanisms of food acquisition such as nematodes have
been shown to readily ingest MPs (Kim, Kim, et al., 2020; Lei et al.,
2018; Shang et al., 2020). In this study we test the following
hypotheses: 1) phagotrophic soil protists ingest MP particles;
2) the number of protists ingesting MPs increases at larger MP
concentrations and; 3) MPs reduce the abundance of active phagotrophic
soil protists.
Methods
Soil collection and preparation
We collected soil from an undisturbed, forested area located within the
Hiawatha Highlands conservation area in the vicinity of the city of
Sault Ste. Marie, ON, Canada (46.5841277, -84.2850883) to reduce the
incidence of any prior MPs contamination. The area is dominated by
Balsam fir (Abies balsamea), White birch (Betula papyrifera) and Sugar
maple (Acer Saccharum). We intentionally included leaf litter and the
top 5 cm of the O horizon because this is where the abundance of
protists is greatest (Adl & Gupta, 2006). We collected 1 kg of field
soil and let it air dry at room temperature over the course of five
days, allowing time for the protists to encyst. Once dry, the soil was
stored in airtight zip-lock bags at 4℃.
Microcosm preparation and experimental design
Six weeks after the soil collection, we placed 10 g of soil in a 10 cm
Petri dish (henceforth microcosm), ensuring that each had similar
amounts of leaf and root material present. We repeated this process for
a total of 30 identical microcosms and randomly divided them into three
treatment groups (n=10). These treatments were prepared as follows:
First, we prepared stock solutions of green fluorescent polymer
microspheres 1-5 µm in diameter (Item # FMG-1.3, density 1.3 g
cm-3, Cospheric LLC, CA, United States) in deionized
water (DI) at two concentrations (1 mg mL-1 (0.1g MP
in 100ml of DI total) and 10 mg mL-1 (1g MP in 100ml
of DI total)). According to the manufacturer, these “highly solvent
resistant” microspheres are made of a thermoset amino formaldehyde
polymer that is inert and fluorescent and is “excellent for PVC and
other plasticizer applications”. Second, we pipetted 10 ml of each
appropriate MP solution (or DI water for the control) into each
respective microcosm treatment for final concentrations of MPs to soil
of 1 mg g-1 (0.1% w/w) and 10 mg g-1 (1% w/w). Despite the scarce
availability of data, these concentrations of MPs may be orders of
magnitude greater than those estimated to occur in the environment
(Jacques & Prosser, 2021). However, this was deliberate to ensure that
we could detect MP ingestion. In addition, the selected size of
microspheres was consistent with the feeding preferences of soil
protozoa (Adl & Gupta, 2006). We considered this the first day of the
experimental trial. We shook the MP stock solutions vigorously while
pipetting to ensure an even distribution of MP beads with each addition.
After adding the MPs, we stirred the soil in each microcosm with a
spatula, to ensure their homogeneous distribution into the soil matrix.
We rinsed the spatula with deionized water in-between microcosms to
avoid introducing MPs. We randomly placed the microcosms in an
incubation chamber set to 22 °C and pre-punctured the lids with a needle
in three spots to ensure proper gas exchange. Microcosms were randomly
placed within the incubation chamber after each time-point assessment of
protists. To confirm the reproducibility of the results we ran a second
trial using the same procedure as in trial one but including the
following treatment groups: no microplastic addition control, and 3 mg
g-1 using the same MPs.
Protist abundance
We quantified protist abundance by direct counts of individual
free-living ciliated and flagellated protists >30 μm in
diameter using the non-flooded Petri dish method (Foissner, 1992). This
involved first adding 10 mL of deionized water to each microcosm to
bring the soil protists out of encystment and counting protists the
following day for a total of 14 days or seven time points (trial 1) and
21 days or nine time points (trial 2). The method recommends eight
sampling points at days 2, 4, 6, 10, 14, 20, 25 and 30. Our sampling
timeline ensured that temporal declines in abundance were captured. More
specifically, the extraction method consists of tilting the microcosm
45° and collecting a small amount of water escaping from the soil. Each
time, we collected six individual 4 μl aliquots (i.e., a total of 24 μl
per microcosm per time point) directly from each microcosm using a
micropipette and placed each aliquot on a microscope slide without a
cover slip. We then observed each aliquot for approximately one minute
using a Leica DM5500B microscope at 50x magnification under phase
contrast microscopy and recorded the total number of protists. We
switched pipettes between samples to avoid cross contamination.
Imaging of MP ingestion
To investigate whether protists can ingest MPs, each time a protist
>30 μm was detected using phase contrast we switched to
fluorescence microscopy (550 nm) and observed it for an additional
minute to look for evidence of MPs (i.e., fluorescent light)
within the food vacuoles. More specifically, if we could see the
fluorescent MPs traveling through the field of view within the living
motile protist’s food vacuoles for at least one minute, we counted that
as evidence of MP ingestion. We also investigated presence/absence of
ingestion in the control group in trial 1 to confirm that other soil
particles or the protist’s organelles would not fluoresce in any
measurable amount. We used the Microscope Software Platform Leica
Application Suite (LAS X) (Leica Microsystems Inc., ON, Canada) to
record video as evidence of MP ingestion while being careful to exclude
any similarly sized organisms other than protists, such as rotifers,
tardigrades and nematodes. Still images were captured from video using
iMovie 10.2.3 (Apple Inc., CA, USA) and processed in Photoshop CC
20.0.10 (Adobe, CA, USA). We conducted this work on days 10, 12, 14 for
all treatments (trial 1) and on days 2 and 6 for the single MP addition
treatment (trial 2). We chose these days to capture a variety of time
periods and to determine if MPs could be readily ingested.
Statistical analysis
The proportion between protists whose vacuoles showed fluorescence
(i.e., evidence of MP ingestion) versus those that did not was compared
between MP addition treatments for each separate trial using repeated
measures ANOVA for days 10, 12 and 14 (trial 1) and days 2 and 6 (trial
2). The negative control treatments were not included in the analysis
because there was no fluorescence detected for any protists in trial 1.
Protist abundance was compared across all three (trial 1) and two (trial
2) treatment groups using repeated measures ANOVA with time and MP
addition as main factors. The data were arcsin (ingestion proportions)
and log (protist abundance) transformed to stabilize the residual
variance. All analyses were conducted using JMP 15.2.1. (SAS Institute
Inc., NC, USA) and data were plotted using DataGraph 4.6.1. (Visual Data
Tools Inc., NC, USA).
Results
Evidence of microplastic ingestion
Evidence of MP ingestion was based on the observation of fluorescence
present within the protists’ food vacuoles (Figure 1 and Figure 2). None of the protists in the control
treatments in trial 1 (a total of 80 individuals across the three
sampling times) showed any evidence of MP ingestion (Table 1). In
contrast, most protists showed signs of MP ingestion in all the
treatments supplied with MPs and this was consistent in both
trials (Table 1). In addition, overall, there was a marginally
significant effect indicating that soil protists tended to ingest more
MPs in the treatment with the highest concentration in trial 1
(F1,9=22.29, P<0.0931) (Table 1). The number
of protists ingesting MPs did not significantly change over time in
trial 1 (F2,8=0.05, P<0.813) and declined from
day 2 to day 6 in trial 2 (F1,9=0.87,
P<0.020).