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Investigating the Effect of Silver Nanoparticles on Aquatic Organisms

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This photo (taken by Eric on December 17, 2007) shows the shell of a ramshorn snail that has been exposed to a silver nanoparticle concentration of 25 PPB.


Since ancient times people have known that silver has antibacterial properties. The ancient Greeks and Romans, for example, put silver coins in their drinking water for this reason. 1 More recently, silver has been found to have antiviral properties as well. 2 Due to its antibacterial and antiviral properties, silver is now being used in an increasing number of commercial products, from washing machines and fabrics to household cleaners and health drinks. 3 Typically, these new uses of silver involve silver in the form of nanoparticles: tiny bits of silver with a diameter in the range of two nanometers. 4 
However intriguing or mundane these new uses of silver might be, an ever greater quantity of silver nanoparticles is finding its way into wastewater and eventually our rivers and streams, where little is known about the possible adverse effects of silver nanoparticles on aquatic ecosystems. In November 2006, the U.S. Environmental Protection Agency decided to begin investigating this very issue. 5 However, it could take as long as 20 years for the EPA to issue regulations for the use and disposal of silver nanoparticles. In the meantime, what effect might silver nanoparticles be having on aquatic organisms and aquatic ecosystems?
I live in Maryland, part of the Chesapeake Bay watershed. I'm very concerned about water-quality issues and threats to life in the Chesapeake Bay and its tributaries. In addition to the widely known pollutants, such as nitrogen and phosphorus, which enter the watershed in huge amounts, I'm particularly interested in the micropollutants that find their way into streams and rivers. For example, trace amounts of estrogens, used in hormone replacement therapy, might be causing male bass in the Potomac River to switch sexes and lay eggs.6 To avoid a repeat of this kind of ecological debacle, I wanted to find out what impact silver nanoparticles might have on the bay and its watershed. I designed a comprehensive experiment to answer my question. 7 

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The simulated aquatic ecosystem: tank A (left), control tank (center), and tank B (right).


The experiment I designed had three parts. In the first part, I tested various concentrations of silver nanoparticles, ranging from 4 parts per million (PPM) down to as little as 27 parts per billion (PPB), on four different aquatic test organisms: Daphnia magna , California blackworms ( Lumbriculus variegatus ), ghost shrimp (Palaemonidae family), and damselfly nymphs (suborder Zygoptera). 8 I chose these four organisms because Daphnia magna is both an example of zooplankton and a common aquatic toxicity test organism; Lumbriculus variegatus is an annelid and another aquatic toxicity test organism; ghost shrimp are crustaceans and aquatic scavengers; and damselfly nymphs are larval insects, important predators in freshwater ecosystems and commonly used indicators of water quality. By using a range of aquatic invertebrates representing different phyla and classes, this part of the experiment gave me an immediate look at the impact of silver nanoparticles on representative organisms in an aquatic ecosystem.

In the second part of my experiment, after I discovered that silver nanoparticles are, in fact, toxic to all four of my test organisms, I tested to see if silver nanoparticles also exhibit secondary toxicity. In other words, do silver nanoparticles continue to be toxic as they move up the food chain, from prey to predator?
In the third part of my experiment, I tested the effect of silver nanoparticles on a simulated aquatic ecosystem. Specifically, I tested two concentrations of silver nanoparticles (that I expected to be sub-lethal) on the growth rate, death rate, reproductive rate, and appearance of five different aquatic organisms: two different aquatic plants Hygrophila polysperma and coontail ( Ceratophyllum demersum ) and three different aquatic invertebrate species ramshorn aquarium snails (Planorbidae family), ghost shrimp (Palaemonidae family), and freshwater clams (Corbiculidae family). Hygrophila polysperma is a hardy rooted aquarium plant, and coontail is a species of SAV, or submerged aquatic vegetation, that's native to the Chesapeake Bay and its watershed. The invertebrates I chose occupy different ecological niches: ramshorn snails are grazing mollusks, ghost shrimp are scavenging crustaceans, and freshwater clams are filter-feeding mollusks. Unlike the in vitro testing in the first two parts of the experiment, this part of the experiment was designed to provide a look at the longer-term, subtler effects, if any, of silver nanoparticles on a freshwater ecosystem. My hypothesis was that silver nanoparticles would be toxic to all of my test organisms at some concentration, and that even at sub-lethal concentrations, silver nanoparticles would have measurable adverse effects on aquatic invertebrates. I also hypothesized that silver nanoparticles would exhibit secondary toxicity.

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Eric adds silver nanoparticles to test tubes.


As it turned out, my hypotheses were mainly correct. In the first part of my experiment—in vitro testing on Daphnia magna , California blackworms, ghost shrimp, and damselfly nymphs—silver nanoparticles at every concentration that I tested, including my lowest concentration of 27 PPB, killed all of my test organisms very rapidly, sometimes in less than two hours. (For details on my procedure and results, please see Appendix 1.) While I expected that at some concentration silver nanoparticles would kill some of my test subjects, the extreme quickness and lethalness of silver nanoparticles was very eye-opening to me. In fact, I was originally only looking for the LC 50 (the concentration that is lethal to 50 percent of the test organisms) of silver nanoparticles forDaphnia , but when the nanoparticles killed all the Daphnia at every concentration, I began to look for increasingly hardy aquatic organisms to test.
The second part of my experiment, testing for secondary toxicity, was nearly as compelling as my in vitro testing. Based on my results, silver nanoparticles do seem to exhibit secondary toxicity. During this part of experiment, which lasted six weeks, I fed ghost shrimp, which are scavengers, either live California blackworms (control) or California blackworms that had died from exposure to silver nanoparticles. (For details on my procedure and results, please see Appendix 2.) Throughout the first several weeks, most of the ghost shrimp remained healthy, but during Week 5, most of the test shrimp started dying and all were dead by Week 6. My control shrimp was the only survivor. The fact that seven of the eight test shrimp died within a very short time frame supports the idea that accumulated secondary toxicity from silver nanoparticles was the cause of death. Unfortunately, this result is extremely bad news for aquatic ecosystems. Secondary toxicity, which has the ability to affect organisms at all stages of the food chain, is perhaps even worse than primary toxicity because secondary toxicity can affect organisms not originally exposed to the pollutant.
The third part of my experiment was the simulated aquatic ecosystem. Unfortunately, the test organisms didn't completely cooperate, as all of my ghost shrimp and some of my clams, including those in the control tank, died. (I think this was because of acidity levels, as the pH dropped below 6, which is fairly acidic for an aquatic ecosystem.) Despite setbacks from the deaths of some of my test subjects, I did get several interesting results that I wasn't at all expecting from this part of my experiment.
First of all, throughout the course of the experiment (four months), silver nanoparticles, even at my lowest concentration of 27 PPB, showed strong algicidal properties, almost completely inhibiting the growth of algae. Given that algae is one of the cornerstones of the aquatic food chain, if my result holds true in an actual environmental setting, silver nanoparticles have the potential to completely change aquatic ecosystems.
Second, although snails laid eggs in all of the aquariums, the number of resulting offspring was lower in the test tanks. (For details on my procedure and results, please see Appendix 3.) The common aquarium snails were the only species of all my test organisms that reproduced successfully. It remains to be seen if silver nanoparticles might inhibit reproduction in other aquatic organisms as well.

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Concern for the health of Chesapeake Bay led Eric to investigate the effect of silver nanoparticles on aquatic ecosystems.


Third, some of the snails in the test tanks developed shell abnormalities, to the point that the shells appeared paper-thin in places. (See Appendix 3 for a photo of a snail shell showing this deformity.) In an environment in which snails are actively preyed upon, this finding could have major implications for snail survival if they can't produce or maintain a strong shell.

All of these results definitely supported my hypothesis that silver nanoparticles would have an adverse impact on aquatic ecosystems. However, my final observation ran somewhat counter to my hypothesis in that silver nanoparticles did not negatively affect growth rates. To the contrary, low levels of silver nanoparticles seemed to promote growth in some of my test organisms, possibly by killing bacteria and algae competing for nutrients and sunlight. This effect might be similar to farm animals that put on weight faster when fed low doses of antibiotics. 9 
Obviously, more research is needed, but my experiment shows that closer attention needs to be paid to the discharge of silver nanoparticles into aquatic ecosystems. Because silver nanoparticles are so remarkably small, it's impossible to filter them out of the water into which they've been discharged. Also, silver nanoparticles seem to bioaccumulate, which means that silver nanoparticle concentrations will continue to increase over time in waterways, including the Chesapeake Bay watershed. Because silver nanoparticles appear to be toxic to a range of aquatic invertebrates, even at extremely low levels, they could profoundly alter our freshwater aquatic ecosystems.
The results of my experiment strongly suggest that silver nanoparticles may indeed pose a serious threat to aquatic ecosystems and their inhabitants. We need to carefully scrutinize our use and disposal of silver nanoparticles.
 

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Daphina Test #1 (September 25, 2007).


Appendix 1: Primary Toxicity Testing Of Silver Nanoparticles In Vitro 
Procedure and Results 
In the first part of my experiment, I wanted to find out if silver nanoparticles are toxic to my representative test organisms: Daphnia magna , California blackworms ( Lumbriculus variegatus ), ghost shrimp (Palaemonidae family), and damselfly nymphs (suborder Zygoptera).
I began with the Daphnia magna , which are small planktonic crustaceans. To do this, I filled sterile test tubes with 10 ml of spring water each. I added 10 Daphnia of approximately the same size to each test tube. Along with the Daphnia , a small amount of their aquarium's spring water also went into the test tubes. To equalize this, I added spring water to each test tube until each test tube had a total water volume of 15 ml. Then I added the silver nanoparticles. The silver nanoparticles that I used were in the form of colloidal silver. This means that the silver nanoparticles are suspended in pure deionized water, with an initial concentration of 20 parts per million (PPM). I added nine different volumes of silver nanoparticles to the test tubes—ranging from 1 drop to 4 ml—and used a negative control that contained spring water and Daphnia but no silver nanoparticles. After I had added the silver nanoparticles, I carefully inverted each tube so that the nanoparticles would be equally dispersed in the water. Every 15 minutes I counted the number of living D. magna. My results are summarized on the next graph.
 

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Daphina Test #2 (October 7, 2007).


Still hoping to find an LC 50 for silver nanoparticles, I repeated the test I did on the Daphnia using California blackworms ( Lumbriculus variegatus) as the test organism. Again, I filled sterile test tubes with 10 ml of spring water each and added a blackworm to each test tube. Along with the blackworms themselves, a small amount of their aquarium water also went into the test tubes. To equalize this, I added spring water until each test tube had a total water volume of 15 ml. Then, I added silver nanoparticles—ranging from 1 drop to 4 ml—to each test tube except one, which was my negative control. I carefully inverted each tube so that the nanoparticles would be equally dispersed in the water. Every 15 minutes, I checked the test tubes to see if the blackworms were still alive. My results are summarized on the next graph.

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Daphina Test #3 (October 17, 2007).


My third in vitro toxicity test tested the effect of silver nanoparticles on ghost shrimp. To do this I began by filling 50 ml-capacity test tubes with 30 ml of spring water. Then I added a ghost shrimp to each test tube, followed by various amounts of silver nanoparticles. In another test tube I had only spring water and a ghost shrimp, making this test tube my negative control. I inverted each test tube to ensure that the silver nanoparticles were dispersed evenly throughout the water. Every 15 minutes, I observed each ghost shrimp and determined if it was living or dead. My results are summarized on the next graph.
 

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Lumbriculus Test #1 (October 19, 2007).


My final in vitro toxicity test used damselfly nymphs. Damselfly nymphs are known to be hardier than some other aquatic invertebrates and able to tolerate "fair" water quality (which is worse than "good" but better than "poor"). To do this, I began by filling 50 ml-capacity test tubes with 30 ml of spring water. Then I added a damselfly nymph to each test tube, followed by various amounts of silver nanoparticles. In another test tube I had only spring water and a damselfly nymph, making this test tube my negative control. I inverted each test tube to ensure that the silver nanoparticles were dispersed evenly throughout the water. Every 15 minutes, I observed each damselfly nymph and determined if it was living or dead. However, the damselfly nymphs were, in fact, hardier than my other test organisms: they did not begin dying until 36 hours after their exposure to the silver nanoparticles. By the end of four weeks, though, all of the damselfly nymphs exposed to silver nanoparticles were dead; the control damselfly nymph remained alive.

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Lumbriculus Test #2 (October 19, 2007).


Appendix 2: Secondary Toxicity Testing Of Silver Nanoparticles 
Procedure and Results 
For the second part of my experiment, I wanted to determine if silver nanoparticles also exhibit secondary toxicity. In other words, do silver nanoparticles continue to be toxic as they move up the food chain, from prey to predator? To do this, I fed ghost shrimp, which are scavengers, either live California blackworms (control) or California blackworms that had died from exposure to silver nanoparticles.

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Lumbriculus Test #3 (October 19, 2007).


I filled 10 new glass bowls with 200 ml of spring water and placed one ghost shrimp in each bowl. Then I waited four days so the ghost shrimp could work up an appetite. Four days later, I placed two California blackworms in each of four sterile test tubes. I added 1 ml of silver nanoparticles to each test tube and inverted it. Once all the worms had died, I transferred the worms, by pipette, to a cup of spring water, to ensure that the silver nanoparticles from the test tubes would not accidentally enter the shrimp water during feeding. Then I carefully pipetted one dead worm from the spring water into each eight of the 10 glass bowls. I pipetted one living blackworm, which had also been transferred to spring water as an intermediary step, to each of the other two glass bowls that served as my control. Within 10 minutes, both the control shrimp and the test shrimp had ingested the blackworms.
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I repeated this procedure, feeding the ghost shrimp every other day for six weeks. Every four or five days, I emptied the bowls and added 200 ml of fresh spring water to prevent the ghost shrimps' water from becoming too fouled. The first feeding was on January 8, 2008.
As the graph below indicates, most of the ghost shrimp remained healthy for the first month of the experiment. Just after four weeks, however, the test shrimp began to die. Seven of the eight test shrimp died within a one-week period, beginning on Day 30, and all were dead within about five weeks of beginning to ingest the California blackworms killed by silver nanoparticles.
Appendix 3: Impact Of Silver Nanoparticles On A Simulated Freshwater Ecosystem 
Procedure and Results 
For the third part of my experiment, I tested the effect of silver nanoparticles on a simulated aquatic ecosystem. After purchasing three new 2.5-gallon glass aquariums, I thoroughly rinsed them out and let them dry completely. Then, using a scale, I weighed out three pounds of Estes Natural Aqua Gravel, rinsed it thoroughly, spread it out evenly along the bottom of the aquarium, and repeated this procedure with the other two aquariums. The next day, I added four liters of spring water to each aquarium. I also set up an air pump with a three-way jack, attached to airstones, to aerate the aquariums and installed a 20-inch fluorescent aquarium lamp that spread light evenly across the three aquariums. I set a timer on the light so that the tanks would receive the same amount of light each day, originally 15 hours per day, to stimulate plant and algae growth.
I decided to add the ecosystem's plants and animals slowly so the nitrogen cycle would have a chance to establish itself. First, I planted 1.1 grams of total biomass of Hygrophila polysperma , a common aquarium plant, to each tank. The next day, I added 1.1 grams of total biomass of ramshorn aquarium snails (Planorbidae family) to each tank. Three days later, I added four ghost shrimp to each aquarium. Two days later, I added 4.6 grams of total biomass of the aquatic plant coontail ( Ceratophyllum demersum ) to each tank. The following day, I added two freshwater clams (Corbiculidae family) to each tank.
Now my simulated ecosystems were ready. I had two types of submerged aquatic vegetation (SAVs), one rooted and one free-floating; two types of mollusks, one a grazer and one a filter feeder; and one scavenging crustacean. Four days later, I added the silver nanoparticles to the tanks. To tank A I added 10 ml of silver nanoparticles, for a silver nanoparticle concentration of 50 PPB. To tank B I added 5 ml of silver nanoparticles, for a concentration of 25 PPB. I didn't add any silver nanoparticles to the third tank because it was my negative control.

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Ramshorn Snails Tests.


Two days later, I changed the light timer from 15 hours to 14 hours per day after I observed large amounts of algae growing in the control tank. However, there was no algae growth in the two tanks containing silver nanoparticles, despite having identical light and temperature conditions. This held true for the four months I ran my simulated ecosystems, suggesting that in addition to having known antiviral and antibacterial properties, silver nanoparticles have strong algicidal properties as well.

Every four weeks for the next four months, I weighed the total biomass of each organism in each tank: Hygrophila polysperma , coontail, freshwater clams, snails, and ghost shrimp. I also checked for snail eggs and observed the snail shells for any deformities.
Because all of my ghost shrimp, both test and control, died early in the experiment, I wasn't able to gather meaningful data from them. Similarly, the freshwater clams also didn't provide me with meaningful data as some of them died (both test and control), and none of them grew at all during the four months of my experiment. Therefore, the only usable data from the simulated aquatic ecosystems came from the ramshorn snails and my two aquatic plants.
 

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Ramshorn Snails Tests.


During the course of my experiment, all three of the test organisms clearly experienced the most growth in Tank B, which had a silver nanoparticle concentration of 0.025 PPM (or 25 PPB). A higher concentration, however, yielded less growth in all cases. This finding would support a conclusion that trace amounts of silver nanoparticles could promote growth in at least some aquatic organisms—perhaps because the known antibacterial or observed algicidal properties of silver nanoparticles reduce competition for resources—but higher amounts may deter growth.
Although egg-laying was observed in all three tanks from the beginning, based on the data in the chart below, ramshorn snails clearly produced more viable offspring in the control tank than in either test tank. It would seem, at least in these simulated aquatic ecosystems, that silver nanoparticles significantly reduced the ability of ramshorn snails to reproduce successfully.
Additionally, some of the ramshorn snails in the test tanks developed whitish patches on their shells where the shell appeared to have been eaten away. The shells seemed very thin in these places (see below). Although not all of the snails in the silver nanoparticle tanks exhibited this sort of deformation, none of the snail shells in the control tank did so.

Endnotes

  1. "The History of Silver as a Bacterial Agent," http://www.agactive.co.uk/information/silvernano/history_of_silver_as_an_antibacterial_agent.cfm
  2. "Study Shows Silver Nanoparticles Attach to HIV-1 Virus," http://www.physorg.com/news7264.html
  3. "This War Against Germs Has a Silver Lining," http://online.wsj.com/article/SB114955908525572199.html
  4. A nanometer is 10-9 meters, or one billionth of a meter.
  5. "EPA to Regulate Nanoparticles Sold as Germ-Killing," http://www.washingtonpost.com/wp-dyn/content/article/2006/11/22/AR2006112201979.html
  6. "'Human Activity' Blamed for Fish Ills," http://www.washingtonpost.com/wp-dyn/content/article/2008/02/07/AR2008020702112.html
  7. The silver nanoparticles that I used in my experiment are in colloidal form, with an initial concentration of 20 PPM, and designed for human consumption. According to the manufacturer, the colloidal silver is nontoxic and no adverse side effects have ever been reported. When I began my experiment, I had no way of knowing whether silver nanoparticles were hazardous to aquatic organisms. Therefore, I was not deliberately exposing aquatic organisms to toxic materials.
  8. All of the aquatic test organisms used in my experiment were purchased from biological supply houses or pet stores. None were collected from the wild.
  9. "Antibiotics as Growth Stimulants for Dairy Cattle: A Review," http://jds.fass.org/cgi/content/abstract/38/10/1102

Bibliography

Blaser, Sabine, Martin Scheringer, Matthew MacLeod, and Konrad Hungerbühler. "Estimation of cumulative aquatic exposure and risk due to silver: Contribution of nanofunctionalized plastics and textiles." Science of the Total Environment 390 (2008): 396-409.
Buzea, Cristina, Ivan Pacheco, and Kevin Robbie. "Nanomaterials and nanoparticles: Sources and toxicity."Biointerphases 2 (2007): 17-71.
Colvin, Vicki. "The potential environmental impact of engineered nanomaterials." Nature Biotechnology 21 (2003): 1166-1169.
Fahrenthold, David. "'Human Activity' Blamed for Fish Ills," The Washington Post, 8 February 2008: B3.
Hamdani, Syeda. Study Shows Silver Nanoparticles Attach to HIV-1 Virus . Physorg.com. Retrieved from the World Wide Web on 12 February 2008. http://www.physorg.com/news7264.html
The History of Silver as a Bacterial Agent . AgActive. Retrieved from the World Wide Web on 8 February 2008. http://www.agactive.co.uk/information/silvernano/history_of_silver_as_an_antibacterial_agent.cfm
Moore, M.N. "Do nanoparticles present ecotoxicological risks for the health of the aquatic environment?"Environment International 32 (2006): 967-976.
Rundle, Rhonda. This War Against Germs Has a Silver Lining . The Wall Street Journal Online . Retrieved from the World Wide Web on 14 February 2008. http://online.wsj.com/article/SB114955908525572199.html
Weiss, Rick. "EPA to Regulate Nanoproducts Sold as Germ-Killing," The Washington Post, 23 November 2006: A1.

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