Figure 13 Location of the Study Area: Samos Island in the Aegean Sea
The research took place near the institute, in the Eastern Aegean Sea, around the island of Samos, close to the coastline of Turkey. The islands’ population is approximately 34000 inhabitants. It has a Mediterranean climate of a typical Greek island, where summers are hot and winters are mild. Furthermore, it has one of the longest periods of sunshine in Greece. (Archipelagos, Institute for Marine Conservation, 2014). The island hosts a high biodiversity, including the surrounding marine environment. The local fishing industry is considered of great economic importance not only for local trade but also for tourism. (Irving, et al., 2014).
The transect lines 1 to 5 show the area of surface sampling near Mesokampo on Samos Island. Most of the fish were caught in Vathi Bay in the North of Island. The second and third S. viridensis samples were taken near Pythagorio in the south east of the island. The fourth and fifth T. mediterraneus were caught in Kokkari Bay also in the north. The fish samples were caught by local fishermen, the information related to the exact location of the catch, such as the longitudinal and latitudinal coordinates are ill-defined due to the lack of GPS information.
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Figure 14 Sample and Catch Location
3.2 Surface Water Methodology
In the following, the methodology to sample surface water and to identify the presence of micro fibres are described.
Surface Water Sampling Method
Sampling took place on the 6th, 9th and 13th of May when the weather conditions were good with low wind speed and sunshine. The method to determine the distribution of microplastic pollution of the surface ocean is described in the following passage. Zooplankton and phytoplankton are both present in the surface water and are prey of B. Boops. Only the presence of microplastic in the surface water is analysed due to the fact that only ingested fluorecent microplastics are visisble under a microscope.
Used materials to carry out this method are:
- Mesh net (a net for sampling the surface water with 333 µm mesh, radius of the haul is 29 cm and detachable collection container) + boom
- Graduated glass sample jars - 5 mm metal sieve
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The first steps of the sampling were to check the net for holes and tears and to note the weather conditions. Afterwards, the boom was used to deploy the mesh net from the side of the vessel to avoid disturbance from the bow wave. The net had to be hauled horizontally through the upper 20 cm of surface water at an average speed of 2.5 knots if possible, over a distance of about 500 meters. It was then dipped slowly down into the water without
submerging the mouth. This helps rinsing the content into the collecting container. Then, the net was retrieved and large debris/ organisms were removed. To collect all debris and plankton stuck, the mesh was washed with seawater in the collecting container. The collecting container was then removed from the net and the sample was reduced to 0.20 litres in a glass jar. After this, the sample was sieved through a 5 mm mesh, so only particles between 0.333 and 5 mm were considered in a jar containing saltwater solution. Finally, the sample jar was labelled and the data such as dates, time, sampling location, length of haul, mesh size (333 µm) and net mouth dimension, were recorded (Archipelagos, Institute for Marine Conservation, 2014).
Laboratory Analysis of the Surface Water Sample
After the field work was properly carried out, the surface water sample was further treated in the laboratory. The required materials are:
- NaCl (33 g)
- Distilled water (200 ml) - Glass fibre filter paper 1.2 µm - Glass syringe
- Vacuum pump and filtration system
The salt solution was made with a salt concentration of 167 g/l. The solution was stirred until the salt dissolved. The salt solution was then filtered twice. To ensure the accuracy and quality of these experiments a control sample was also produced.
After the sample was sieved into the jar (see surface water sampling method), the sample was left to settle in the glass jar for 24 hours to separate the low-density plastic particles from organic tissue by buoyancy. After settling, a glass syringe held at the sample surface was used to extract the supernatant. This extracted water was then filtered; the retained microplastics were washed with distilled water to remove the salts. Thereupon, the filter paper could dry it was covered while drying to avoid contamination. The microplastic fibres present were expressed in particles per fitered volume (particles/m3). The filtered volume is described with the following formula. 𝐹𝑖𝑙𝑡𝑒𝑟𝑒𝑑 𝑉𝑜𝑙𝑢𝑚𝑒𝑛 = 𝜋 ∗ 𝑟2∗ 𝑑. Wheras, r is 14,5 cm (Archipelagos, Institute for Marine Conservation, 2014).
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3.3 Fish Stomach and Gut Methodology
The fish stomach and gut methodology not only consists of the dissection of species.
It also includes the analysis of microplastic fibre present in the stomachs of mentioned species. Each individual sample was weighed, measured, dissected and the sex was determined. To ensure reliable results, five stomachs were analysed for each trophic level.
The following dissection description gives an idea on how the stomach was identified and removed. The stomach and gut of each fish was used to determine the microplastic fibres present in the individual.
Dissection Description
This method gives an overview of how the dissection was carried out. The materials used for fish dissections are:
- Dissection microscope with a proper illumination - Dissection set with scissors, scalpel and forceps - Probes (needle with blunt tip)
- Kitchen bakery paper
- Lab coat, gloves (non-plastic) - Security goggles
- Glass Petri dishes.
As shown in figure 15 “Incision with a Sharp Knife”, the first step of a fish dissection, is to make a shallow longitude incision along the ventral left-hand side which extends from the anterior of the anus to below the gill arches. It is important to cut deep enough to gain access to the gut but damaging the organs has to be avoided (Elenbaas, 2014).
Figure 15 Incisions With a Sharp Knife (Elenbaas, 2014)
Afterwards, the hand is put inside the gut cavity and one side of the cage is gently lifted up.
Then, transverse cuts, from the anterior and posterior ends of the longitudinal incision on the
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left side of the specimen are made. It is valid that the transverse cuts extend as far dorsally as required to permit an unobstructed view of the gut cavity. At the end, the stomach and gut is identified (see Figure 16), removed and placed in a Petri glass (Elenbaas, 2014).
Figure 16 Overview Organs in a Fish (Archipelagos, Institute for Marine Conservation, 2014)
Analysis Method for Microplastic in Fish Stomach
As soon as the fish stomachs were removed, the microplastic contamination was evaluated. It was important to clean the materials prior with alcohol and three times with filtered distilled water. The required materials were:
- NaCl
- Distilled water (Cole, et al., 2014) - Graduated glass jars with metal caps - Glass fibres filter paper 1.2 µm - Forceps
- Tape measure scale (Archipelagos, Institute for Marine Conservation, 2014)
The salt solution was made with a salt concentration of 167 g/l. The solution was stirred until the salt was dissolved. The salt solution was then filtered twice. To ensure the accuracy and quality of these experiments a control sample was also produced. The stomach samples were placed into the saltwater solution. The glass jar was shaken for one minute and then left for about 24 hours, so the organic material could sink and the low density plastics float on the surface of the solution. Afterwards, 300 ml of the supernatant was removed with a glass pipette at the surface. This extracted water was filtered and the filter paper was allowed to dry.
The jar was filled with filtered salt solution, again shaken and left for 24 hours, to ensure that
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as much microplastic as possible was collected. Once more about 300 ml of the supernatant was removed and filtered. To dry the filter paper, it was placed in a glass petri dish and covered to avoid contamination (Archipelagos, Institute for Marine Conservation, 2014).
3.4 Filtration Paper Analysis
The filtration paper method was used in both cases, for the analysis of microplastic contamination for fish stomachs and zooplankton.
Figure 17 Method to Review a Filter Paper (Archipelagos, Institute for Marine Conservation, 2014)
After vacuum filtration, the clamp and top piece of the Whatman flask needed to be removed.
The filter paper was transferred into a glass Petri dish by using metal forceps. It was
important not to disturb the sample at any time. For analysis, the filter paper was put onto a glass slide using, again, metal forceps. The slide was then placed under the microscope.
Using a 40x magnification lens the filter paper was systematically viewed, using the method shown in figure “Method to Review a Filter Paper”. During this viewing items of interest were counted and, to ensure accuracy, a second count was carried out (Archipelagos, Institute for Marine Conservation, 2014).
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3.5 Method Review
To prevent contamination during lab work, the distilled water from a plastic container, used to prepare the saltwater solution, was filtered before use. Additionally, the equipment was checked for contamination under a microscope and surface, hands and equipment are cleaned with distilled water before use. During the dissection, nitrile gloves and not plastic ones were worn to prevent manipulation and possible contamination of the stomach with microplastic fibres (Archipelagos, Institute for Marine Conservation, 2014).
The literature research showed different method to dissolve organic matter (described in chapter 4.1). Due to the lack of time, further research on new methods to dissolve a fish stomach and the correct ratio of baking soda to water, stirring and temperature was not able to be tried out. Additionally, the used stomach to test the methods was of a B. boops, it is significantly smaller than one of larger fish, the barracuda and mackerel. The time needed to dissolve the larger stomachs is therefore longer and needs further investigation. Another reason to not continue investigating different methods was that the laboratory equipment did not allow for proper contamination control. The stirring magnet was covered in plastic which might erode and skew the results; the temperature measuring device needed to be placed in the solution whilst heating up; and the baking soda is packed in plastic, all of which may lead to contamination. Other ideas, such as the usage of an ultrasonic bath to dissolve the
organic tissue could not be conducted due to the lack of equipment.
Ultimately the saltwater method, mentioned in various other researches and previously used by Archipelagos, was used to investigate the presence of microplastic fibres in the fish stomachs. Reviewing several reports on microplastic presence in plankton and fish gut showed that the techniques which are used for this research are quite uniform. Similar methods are described in “Microplastics in the Marine Environment: A Review of the
methods Used for Identification and Quantification” ,by Valeria Hidalgo-Ruz et al., “Isolation of microplastic in biota-rich seawater samples and marine organisms” ,by Matthew Cole et al., “The Impact of Plastic debris on Biota of tidal Flats in Ambon Bay”, by Prulley Uneputty and S. M. Evans, and “ Plastic ingestion by planktivorous fishes in the North Pacific Central Gyre”, by Christiana M. Boerger et al.. One disadvantage of this method is that it only enables the detection of low density microplastic fibres, high density fibres cannot be recorded.
Another problem may arise while using a 0.02 µm filter paper, as clogging of the filter may occur due to improperly dissolved tissue and salt crystals. A mesh filter of 50 µm appeared to have the fastest filtration rate. Although the 20 µm mesh-filter theoretically captures smaller microplastic nevertheless those small particles are able to be identified with only a binocular
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microscope. It was therefore not necessary to use such a small mesh size for this study (Cole, et al., 2014).
3.6 Statistical Analysis
The statistical analysis is carried out by determining the standard deviation for weight, length, number of fibres, number of fibres per weight or length. The following formula is used to determine the standard deviation:
𝜎 = √ 1
𝑛 − 1∑(𝑥𝑖− 𝜇)2
𝑛
𝑖=1
Furthermore, the correlation coefficient r is determined by using a power trend line. The correlation coefficient gives an idea about how strongly related one variable to another is. It ranges from - 1,00 to + 1,00. A perfect negative relationship exists between two variables when r is - 1,00. On the other hand, a perfect positive relationship occurs whenever r is + 1,00. The closer r is to - 1,00 and + 1,00 the more correlation exists (Higgins, 2005).
A power trend line is curved and preferably used with a data set that compares
measurements which increase at a specific rate. It can only be used whenever the data contains no zero or negative values (Office, 2015). This is the case with the data set
collected during this study, due to that fact that no fish in existance weights or measures zero or negative weight or size.
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3.7 Knowledge Gap, Limits and Preconditions
It is unknown at what time microplastic started to be present in the sample species.
Another knowledge gap is the extent of microplastic content in the target species of this research. Currently very little research is being carried out with the species being
investigated. Furthermore, it is not possible to clarify if the species ingested microplastic mistakenly, on purpose, or indirectly through their prey. Moreover, pollution might be a result from other prey species and not only the investigated one; microplastic pollution could also be a result of preying on M. surmuletus and not T .mediterraneus. In addition, it is not possible to have exactly the same location for all the samples. It is only possible to have samples from the same region so there might be discrepancies in their contamination with microplastic due to differences in living conditions and surrounding.
The research was limited by weather conditions which influenced the possibility of obtaining fish and taking the surface water samples. The surface water samples are only able to be taken with low wind and wave conditions.
A sodium hydroxide solution with 10 M would be a suitable method to decompose all organic matter, but it potentially leads to damage or discolouring of the plastic particles can occur with such a strong alkaline treatment (Cole, et al., 2014). Further, the lack of a laboratory hood makes it too unsafe to use a sodium hydroxide solution with a concentration of 10 M or at a temperature of 60°C, due to formation of toxic gases (Cole, et al., 2014). As a laboratory hood is not available, neither attempting the sodium hydroxide method nor the potassium hydroxide method are options for this research.
The basic laboratory equipment did not allow any other strong alkaline methods or acidic solutions. Furthermore, the lack of a tap in the laboratory makes it difficult to maintain a clean working environment. Additionally, the microscope can only identify plastic particles which are big enough to be identified with a 40x magnification. This also only allows the
identification of colour and existence of microplastic. The size of the fibres is not able to be determined because they are not present linearly and this cannot be measured. It is also not possible to analyse the type of plastic due to insufficient equipment.
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