We Have Numbers Of Free Samples

Abstract

Freshwater ecosystems can be easily and unknowingly polluted by the surroundings, this can cause detrimental damage to the ecosystem.  Mayflies are a biological indicator and are an effective yet simple way of measuring an ecosystem’s health. I collected data on the distribution of Flattened Mayfly Nymphs in order to further inform other related ecological studies. I used a kick sampling method at multiple areas to collect 15 samples and measuring the flow velocity using an impellor and hydro prop at every site. My final results showed a positive correlation of abundance to flow velocity of  (dA/dV = 15.62) and using a spearman’s rank test found there is a 99.5% chance my data has true correlation.

Research and Rationale

May Fly Life Cycle (3)

 

The mayfly belong the one of the oldest order of insects, the Ephemoptera with an evolutionary history dating back to up to 300 million years. There are up to 50 species of Ephemoptera in the UK, I will be focusing on the genus Ecdyonurus however I am unsure of the species as it is difficult to identify the four main UK species of Ecdyonurus nymphs. The Flattened mayfly nymph does not tolerate polluted water so its presence indicates a relatively pristine habitat, making them a good biological indicator for mesotrophic conditions (3). They are considered an ecological keystone species linking detrital energy resources directly to higher trophic levels containing many species of fish which feed on the nymphs(4). Meaning the expiration in an area of Mayflies could lead to ecological collapse(4). Mayflies are useful indicators as they are relatively easy to sample, highly visible and provide evidence of effective water quality control. I will be basing my study on them as the implications of monitoring their distribution and abundance allows the ability to further inform studies regarding Flattened Mayfly Nymphs and pollution.

The life of a mayfly begins once it’s hatched from it’s egg, spending up to 2 years in this Nymph form at the river bed feeding on algae and detritus, they have a very hydrodynamic form and relatively large tarsal claws, the combination of the two makes them better adapted to fast flowing currents and are found clinging to rocks or algae(1).

After their nymph stage is finished, at about June or October time they rise to the surface and break through the surface tension. Once there, they shed their nymphal skin becoming an “emerger” until their wings dry and they take flight, they are then considered a “dun” as they are not able to reproduce. The dun will then undergo its second metamorphosis becoming a spinner where it’s only purpose it to mate. The females then dip in and out of the water dropping it’s eggs off, these are named “Ovipositing spinners”(2).

The name of the order Ephemoptera is derived from “ephemeros” meaning literally “lasting a day”, this is because their adult stage lasts only a day. From their first metamorphosis they lose their digestive tracts and are replaced with their sex organs and they have no functioning mouth parts. Their entire adult stage is limited by their energy reserves created as a nymph hence their short adult life span(1).

Flattened mayfly nymphs prefer stony substrata such as riffles; this is because of the faster flowing current and larger stony substrata where they are better adapted to, allowing them to cling onto rocks also hiding themselves from predators. A study published in the Canadian Journal of Zoology in 1997 explored the effect of flow velocity and light on mayfly nymphs, the result showed a preference to faster flowing darker waters(5). They have obvious adaptations allowing them to inhabit faster flowing water unlike other organisms in a similar habitat which dwell in slower currents.

Tarsal Claw of Heptageniidae(1)

 

The adaptations which lead them to preferring riffles include their tarsal claws which are very pronounced allowing them to cling on to stones and their flat hydrodynamic form reducing the resistance from the current, both of which stop them being swept downstream⁽¹⁾. They have a very dark/transparent camouflage reducing the chance of preditation especially when they are relatively idle when clinging.  All of these adaptations expand their niche to faster shallower areas such as riffles. This presents the advantages of a reduction in competition for nutrition and space.

Another study conducted in Oregon published 1984 explored the effect of substrate type on distribution of deferent species of Mayfly⁽⁶⁾. The samples present data from large stony substrata to small sediment (<1mm) also containing data on moss and algae’s effect. This study shows me that Substrata and moss growth do have an effect on the Mayfly nymphs distribution and clarifies that different species’ abundance is affected differently. This study also shows flow velocity and sediment type are correlated as only larger heavier rocks are able to remain in higher currents. Relating back to the Canadian Study above.

Food availability is also another factor, the above study shows a slight influence of Algae and Moss on certain species of mayfly. I know the flattened mayfly nymph feeds on Algea, moss and detritus, however in faster flowing currents, where the studies suggest a higher concentration in distribution of flattened mayflies, there will be a lack detritus so I can say that Algae and Moss will affect their distribution. As they’re both photosynthesizing I can therefore predict that light will also have an effect on their distribution.

Taking all into account I can come to the following Hypothesis:

Hypothesis

H¹: There will be a positive correlation between the velocities of the site on the river.

Planning

Pilot

For my pilot test I need to measure my confounding variables to ensure they do not affect the distribution of Flattened Mayfly Nymphs and test my biotic and abiotic collection methods for the most effective and efficient methods.

In my pilot I will find the following:

• The most efficient method for biotic sampling, kick sampling or Dragnet sampling.
• Measure the confounding variables, Temperature, PH and dissolved O₂.
• The most appropriate method of collecting flow velocity.

To do the above I will use the following equipment:

• For the Biotic sampling I require a D framed net, tray, bucket, spoon and wide opening pipette.
• For my Velocity measurements I will require an impellor and hydro prop to compare to the float method.
• Electronic PH metre and a universal PH indicator kit.
• Electric thermometer.
• A dissolved oxygen metre.

Risk Assessment

Risk	Step Taken To Reduce Risk
Slipping on rocks/muddy surface	Wearing appropriate foot wear
Drowning	Keep in couples and being cautious
Sun Burn	Sun lotion and a hat
Hypothermia	Appropriate warm clothing
Trampling by cows	Keeping a safe distance
Biological Hazards	Washing hands on return
Working alone	Do not work alone

 

Potential Hazards	Worst Case Scenario					Probability
	Minor Injury (1)	Injury (2)	Major Injury (3)	Crippling Injury (4)	Fatality (5)	Very Rarely (1)	Rarely (2)	Infrequently (3)	Sometimes (4)	Often (5)	Risk Rating
Slipping on rocks/muddy surface		/						/			6
Drowning					/	/					5
Sun Burn	/								/		4
Hypothermia		/					/				4
Trampling by cows				/		/					4
Biological Hazards		/						/			6
Working alone				/		/					4

 

Risk Rating = Worst Case Scenario * Probability

My total risk is 33 out of a total of 175, I consider this relatively low risk and conclude my experiment is safe enough to continue.

Method

I will collect biotic data working upstream, from pools and riffles as a method to represent velocity of the stream; this will allow me to find if there is a correlation between the velocity and the abundance of Flattened Mayfly Nymphs.

In order to decide between my two collecting methods I will kick sample in a pool and riffle for 10/20/60 seconds then I will drag sample for 0.5/1.0/1.5 metres. This will allow to compare the methods against each other and the most efficient time of disturbance or distance dragged. This will give me a total of 12 biotic readings. I will measure the streams velocity in the same place using both an impellor and a float to give me an idea on the best method to use for my main investigation. I will do the same with the electronic PH metre and the universal indicator.

The temperature and dissolved oxygen levels will be measured throughout the stream to find if they fluctuate and therefore be measure in my main investigation.

Review

For the biotic methods, I found that the dragnet method didn’t work in the pools as it was impossible to count the mayflies as it brought up allot of sediment, leaves and rocks.  On average the kick sample method found more Mayflies per minute than the dragnet method. So I will use the kick sample method on the main investigations as it is more efficient and effective.

Also, the biotic readings showed me on the kick samples performed for the same time, there were more Flattened Mayfly Nymphs in higher velocity waters (Pools) compared to slower flowing water (Riffles) thus showing plausibility to my hypothesis leading me on a good path towards my main investigation.

There was no significant change in the P.H from what I could tell; I used both universal indicator and an electronic P.H. metre. The P.H. appeared different on the universal indicator to the electronic P.H. metre by about 0.8, this was because first the indicator was inaccurate, it only found it to the nearest X.5, considering the electronic metre’s value in between 7.5 and 8.5 it explains why the indicator was far out of the true value. Also the indicator was a lot more subjective than the electronic metre allowing it to be open to interpretation where as the electronic metre gave an exact value. Although the two methods results were quite far off each other in their measurement it still told me that the P.H. was a constant throughout the river and therefore it will not be measured in my main investigation.

The temperature did not change by a considerable amount, I found it be 0.2 oC lower in the pools however I do not consider it to be large enough to affect my results and therefore I will not measure this in my main investigation.

While conducting my dissolved oxygen I found the metre to be very unstable, dropping in its value for about five minutes or longer then rising, sometimes varying by as much as 1mg/l due to clouds sediment interfering with its measurements as it uses the conductance of water to determine the dissolved Oxygen. Once I found them to finally stabilise they were still able to vary by about 0.1mg/l in both directions. The measurements I was able to attain did not show a considerable difference between the pools and riffle’s. I therefore do not consider the Oxygen worthy of measuring as I did not find any considerable change and combined with the fact it takes a long time and is unreliable I do not believe it is worthy of measuring in my main investigation.

Summary

I found the following from my pilot:

• I will use the kick sample method
• I will not measure the P.H
• I will not measure the temperature
• I will not measure the dissolved oxygen levels
• I will measure the velocity and abundance

Main Investigation

Variables
Independent Variable : Velocity of stream
Dependent Variable: Abundance of flattened Mayfly Nymphs
Confounding Variables: Shading of area (Mayfly Nymphs feed on photosynthesising algae)
Temperature, Depth, PH
Biotic factors (such as different forms and abundance of prey)

Method

Sites

I will work up stream aiming to collect data for both biotic and abiotic from 30 sites (as that is my data test’s limit) along a stretch of the meander near Nettlecombe Court, thus allowing a certain degree of control over my confounding variables in the sense I can assume: Temperature (which I measured to be relatively constant in my pilot study with a maximum variation of 0.6 C), PH (which I also measured to be constant with no variation in my pilot), Dissolved Oxygen content (which I measure to have a maximum variation of 0.2 mg/l ) and Preditation (as it is a single micro-ecosystem so to have little variation) all to be constant for the purposes of this study throughout my selected stretch of meander. I will select my sites will be in similar lighted areas as to further control my confounding variables, I will do this by avoiding heavily shaded areas. I will work from pools to riffles collecting 3 or so data sets along the way; this will hopefully give me a wide range of values for velocity.

Biotic

For collection of my biotic variables I will require the following equipment:

• Light coloured tray
• D-Frame net
• Bucket
• Wide nosed pipette
• Stop watch
• Plastic spoon
• Magnifying glass

1. I will select an appropriate site not shaded by trees so to affect the light in this area and no one working upstream so their disturbances could not affect my results.
2. I will first fill my tray and bucket with water from the stream, so I could keep them in the tray and bucket to count without causing harm to them, being careful not to disturb the floor of the river so I do not unsettle them.
3. I will then carefully walk into the stream with the D-Framed net, placing it onto the floor, starting the stopwatch and beginning disturbance upstream from it with my foot moving in a circular motion applying a fair amount of pressure, being careful not to kick the rocks into the net. I will stop after 20 seconds as my pilot showed this to be the most efficient time.
4.  Once 20 seconds has passed I will lift the net out the river whilst pulling it towards me so nothing escapes. I will then invert the net into the tray and use the bucket of water to rinse the net ensuring everything is inside the tray, a light coloured tray makes the dark Flattened Mayfly Nymphs easier to spot.
5. I will remove any large sediment from the tray, shaking it off in the tray submerged thoroughly so any clinging nymphs are left inside. This makes it easier to count them as they cannot hide beneath them. I will return the sediment to where I found it so not to disturb their natural environment.
6. I will then measure the abiotic factors (I will explain this bellow), this allows the sediment to settle whilst I measure the abiotic so on my return it will be easier to count them and so I don’t have time to forget exactly where it was I took my measurement.
7. I will then count the number of flattened mayfly nymphs inside the tray. I will do this by first sucking them up into the wide nosed pipette and emptying it into the plastic spoon allowing me to clarify if necessary with the magnifying glass that it is a Flattened Mayfly Nymph. Once I am sure I will then put it into the bucket so to avoid double counting an individual also stopping to record every 5 I find making it more efficient. Once there are no more in sight, I will use the pipette to disturb any stones or sediment in the tray to get any out of hiding.
8. I will spend a maximum of ten minutes counting so not to waste time. Once I am done at the sight I will empty the tray and bucket out back into the meander near to where I took them. Giving the bucket and tray a rinse to ensure they’re all out and moving on upstream to my next site, repeating this process.

Abiotic

For collection of my abiotic variables I will require the following equipment:

• Impellor
• Hydro prop
• Stopwatch
• Half metre rule

1. It is essential the abiotic readings are taken after I have collected my sample to I do not disturb them, possible causing anomalies in my biotic data.
2. I will first measure the velocity of the stream at the site by timing how long it takes the impellor to travel the length of the screw in seconds, I will do this by stepping into the river, standing behind the impellor so not to affect the flow, setting the impellor and then holding it down with the hydro prop on the floor starting the stop watch at the instant it is submerged. Once the impellor has reached the end I will stop the stop watch. Without moving out the river for efficiency and reliability I will reset the impellor and note the time. I repeat this three times in the same spot for as I found the velocity measure can vary in the same spot during my pilot and I will stay in the river so to cause minimal disturbance. My recorded impellor time can later be converted into ms^-1 later using the following formula:
3. I will then collect my depth data in metres using the half metre rule. I will hold the rule down to the bottom with its thinnest side facing the current so the water does not wrap around it giving me a false value. I will take four measurements in a square configuration avoiding the ditch creating by my disturbance.

Site Map

Bellow is a site map of the meanders showing where I collected my data. I worked upstream from piont

Site Map (courtesy of Google Maps)

 

Abundance of Flattened Mayfly Nymphs Throughout The Stream

Evaluation of Sources

(1) This article is on the APEM website. APEM is Europe’s leading environmental consultancy specialising in freshwater and marine ecology and aerial surveys. It is written by two scientists both with long careers in fresh water ecosystem and Ph.D’s based in similar areas. This being an independent company and who reputation relies on sound scientific studies, I do not consider this to be a bias source and I also onsider it reliable.

 (2) This is website named “digital Key To Aquatic Insects” set up by the macro-invertebrate lab in Valley City University, North Dakota. This specific article is written by Dr Andre DeLorme, their associate Professor of Biology, he earned his Ph.D in Entomology from the University of Minnesota. He has released papers on the effect of pollution on fresh water shrimps and many others. With no bias, especially to information about mayflies, and his qualification I consider this a reliable source.

(3) This is the source of my picture on the Mayfly’s life cycle, it is made by “Timsbury fishery” as an informative handout for fisherman. This is not a particularly reliable source however for its purpose of the picture I do not consider it an issue as it agrees with the information from my other sources.

(4) This is an article written about Burrowing mayfly nymphs. It is on the United States government Environmental Protection Agency‘s site as an informative piece. I consider this a highly reliable source as its sole intention is in the protection of the environment

(5) This is an in depth study concerning mayflies, I read the article at Sussex University Library. It has 8 citations in many reputable journals such as “Ecography” according to Google scholar. The citations are by well educated Doctors. There is no obvious bias towards this study and for this to have been published in this journal it would have been subject to peer review, also I consider this studies methods to be scientifically sound. Taking all into account I trust this study.

(6) This is a study published by the Professor of Aquatic Ecology & Watershed Sciences at Utah State University.  He is a very active ecologist having involvement in many studies; he has had 2939 citations since 2009 according to Google Scholar. I consider it a reliable source and I personally consider his methods appropriate and reliable. Also to be published in this journal it would have been subject to thorough peer review.

(7) Michel Sartori is the professor of Zoology at the university of Hamburg. He has had a total of 1032 citations according to Google Scholar. The journal of Aquatic Sciences is a well known and respected Journal and their methods are appropriate and have been subject to peer review. Therefore I consider this a reliable source.

(8) This is the federal environmental protection agency in the US. They monitor the environment and it’s potential effects on human health. Being a government run agency I can assume it’s free to bias and as many respected scientists behind them. This is an instructional article on how to perform reliable stream biodiversity studies.

(9)This a masters student’s site on aquatic invertebrates with images and information. There is no sensitive data here venerable to bias, just some brief information about nymphs and many close up pictures.

[Download not found]

Download