|This chapter instructs you how to do the DDA test and how to prevent common mistakes. Because you are now doing science, you must adopt precise habits and constantly be aware of possible mistakes. The DDA is by scientific standards a very simple test, which junior college students should be able to do successfully. Read this manual carefully and keep it with your test set for easy reference. Be aware that this chapter on Internet is updated from time to time as new developments come to hand.|
|A short introduction to the DDA method. For a more complete introduction, please read the DDA index page which also has a dictionary of strange and new words.|
|Although it looks unassuming, the pH meter is a technological achievement of precision that needs to be cared for and calibrated often. Your results depend on it entirely. See also the document ph.htm explaining more.|
|Scientists have access to a large assortment of containers to be used as vials but we took vials that are readily available from your photographer: film cans.|
|A 50m long non-stretching measuring tape as used by surveyors, enables you to measure the visibility of the water you are testing. Here are some tips and tricks.|
|When measurements are done at a constant temperature, they can be compared whether taken by different people, at different times or places. Your travel incubator is a special instrument.|
|Not only temperature must be standardised, but also the sequence in which measurements are done, and how many in total.|
|How to calculate biodensity and rate of attack.|
|How to dilute the ethyl alcohol to a standard solution of 20%|
|How to prepare your used vials for a new test.|
|Here are some useful documents and images:
While searching to explain the degradation occurring in the sea, Dr Floor Anthoni discovered that the planktonic decomposers had been overlooked as a most important missing ecological factor (limiting factor). That enabled him to discover a method to measure not only the activity of the decomposers but also the total decomposable biomass, which could consist of plant plankton, animal plankton, bacteria and other biological material such as raw sewage and undefined dissolved organic matter (DOM). It is important to understand that the Dark Decay Assay (DDA) is not able to tell these apart.
The DDA is indeed quite simple in principle: take a water sample in a bottle or jar. Shake it so that it becomes evenly mixed. Pour it through a coars sieve (strainer) into containers (vials) that can be sealed, and fill these as full as is possible. Screw or press the lids back on and store the vials in a perfectly dark place at a constant temperature. About once a day, open each container and measure its acidity (pH) using an accurate three-digit pH meter. Plot the results on graph paper or on computer. After one week at elevated temperature which accelerates the decomposition, all biological matter has been decomposed. Wait another week to be sure.
What happens inside the vials? Because plants cannot survive in the dark, the plant plankton immediately stops working and eventually dies, which happens surprisingly quickly, already after two days in the dark. The decomposers which consist of single-celled bacteria, fungi and viruses, will decompose the plant plankton and eventually also one another. Because they break apart the biochemicals of life which consist of carbon chains with many hydrogen bonds, hydrogen ions (H+) are returned to the water which makes the water more acidic. Because the lids are air tight, these hydrogen ions cannot escape in the form of gases, and they are measured by the pH meter. Thus the pH is an indication of the amount of biomass decomposed, and this can be plotted as curves on a graph.
The decomposition happens more quickly at higher temperatures, which means that the shapes of the curves depend on temperature. If it gets colder, the curve flattens. If it becomes warmer, the curve steepens. So it is important to stabilise the temperature for the following reasons:
minimum DDA laboratory set contains the following items:
The DDA method works only if all vials are kept in the dark at all times. The incubator should therefore always be securely closed as it also functions as a dark box. But when samples are measured, they are also exposed to the light, which is to be minimised:
As part of the measuring routine, it pays to shake each container a
few times in order to stir its contents, but we have not been able to demonstrate
that this actually matters.
Although the water looks clearer than drinking water (which it is most times) you may forget that inside lives an ecosystem containing millions of individuals of thousands of species. It is a true mini world to such an extent that each vial decomposes in a different way compared to every other vial. Thus even when samples are taken from a well mixed jar, differences between them will develop. So it is a good idea to take more than one sample from an evenly mixed jar. For non-scientific work about two samples are adequate, but for more precision and to satisfy scientific criticism, more samples are required from a jar taken at one place at a given moment in time. This of course can easily double the work later. One can often limit the number of duplicates when a batch consists of mostly similar samples taken from similar bodies of water, following a coastline for instance.
Having many vials with many different microbes inside them, raises the possibility of cross-contamination. In its worst case, you will end up with each vial containing the same stuff as all the others. Thus minimising cross-contamination should also be a priority. Ideally one would like to sterilise the pH probe each time before inserting it into the next vial but this raises other problems. Fortunately for us, our samples behave more or less like fully populated communities where introduced strangers are not noticeable and cannot multiply unless there is vacant space available. Thus cross-contamination is fortunately not very serious.
But there's more to it. Where we measure the Rate of Attack in the first 48 hours, cross-contamination is still so minimal that it does not play a role. The beginning of each decay curve is truly due to the decomposers already in the sample and to those that are already numerous. What happens later, is less important. But we noticed that complete decomposition is often not happening unless a small amount of cross-contamination has occurred. It seems as if all the bacterium species necessary to decompose other bacteria, must somehow be present and if not, must be introduced in very small numbers.
To minimise cross-contamination, always proceed as follows:
In order to be successful, you must commit yourself to your experiment such that the measurements are done timely and accurately, a spying eye is kept on things that could possibly go wrong, and things are kept tidy. One of the most important tidiness should be the way in which you record your results. Write clearly such that there can never be doubt about the figures. In case of mistakes, use a typist's white-out method so that in the end, the data cards look tidy and anyone can clearly read and transcribe them. Remember these cards carry the results of all your work. Make it a good habit to do backups now and then by photocopying all cards that are ready.
All items used have a name like A1, A2, B1 or ALPHA, BRAVO and so on. Where more than one pH meter is used, they also have unique names or numbers. All this is necessary in order to be able to identify the sets used, without confusion so that irregularities can be tracked down. For instance, a certain vial may always be out of line, possibly caused by a crack in the material. This can be tracked back because all results have been labelled.
Once an experiment has completed its course, the data must be plotted to make sense. Plotting cards are provided and a four-colour ballpen. This will need some experience.
|The pH meter
Although the portable battery-operated three-digit pH meter looks unimpressive, it is nonetheless a technological achievement of high precision. Your results depend on it entirely, so it needs to be looked after and recalibrated often.
Although a new pH meter is delivered completely dry, it is not advisable to let your pH meter's electrodes dry up. It would eventually build a crust of minerals that could affect its precision. A new pH meter must be soaked first in a saline solution, for which a vial of potassium chloride (KCl) is provided. After an hour the pH meter can be calibrated. How this is done precisely, is found in the meter's manual. But it goes in principle like this:
You have been provided buffer solutions with known acidity or alkalinity:
pH=4.01, 7.00 and 10.01. Chemical buffers have the property that their
pH values are not easily disturbed by contamination, but we will attempt
to minimise this. Your pH meter assumes that when it is presented a solution
close to pH=4, and the calibration button is pressed, that the solution
is exactly pH=4.01, and it will make an internal adjustment accordingly.
Begin with pH=4 because it has the most hydrogen ions.
Rinse the pH probe with tap water, shake it dry and present the pH=7.00 solution. Once the reading has stabilised, which can take ten minutes, press the necessary buttons to calibrate. Finally the pH=10.01 solution is done.
Because our measurements range from pH=6.2 to 8.2, the middle value pH=7.00 is very important whereas the furthest away, pH=4.0 and pH=10 are least important. Thus the pH=7.00 calibration is done most often and with most accuracy. That is why you keep three vials with pH=7.00 buffers. One is labelled FIRST to take any contamination first. The other is least contaminated and most precise. Then there is also a spare just in case. You can get new pH buffers from a friendly laboratory or from a laboratory supplier. They recommend that buffers be replaced every year, as they have an expiry date. You can extend the life of buffers by storing them in a dark, cool place.
When a pH meter is new, it drifts so much that it needs to be recalibrated at least twice daily. After a few days it becomes more stable and after a few months it may need calibration twice weekly. Make a note of the old pH=7 values before pressing the calibration buttons. When a pH probe is faulty, it can be replaced without replacing the body holding the electronics. Every time a probe is recalibrated, it is registered in your calibration log. This allows you to detect problems at an early stage.
Scientists have access to a large assortment of containers to be used as vials but we took vials that are readily available from your photographer: HDPE (high density poly ethylene) film cans. These HDPE cans protect the sensitive photographic emulsion from dust and moisture, so they should also be air tight. Unfortunately this is not securely the case for most brands, and we tested the FUJI film canisters with transparent bodies and transparent or black lids as the best choice. Their lids push down INSIDE the container into a polished groove. Test the lid's fit for each container, by rotating it.
lids of these canisters are shaped such that a bubble can be formed right
in the top of the lid, allowing the lid to be pressed close. With a cork-borer
tool, a hole can be cut in a lid such that the pH probe fits securely in
the hole of the collar, and then onto the vial. It fits sufficiently neatly
that the container can safely be lifted with the probe. The probe needs
to protrude only a little, as the liquid pushes up around it on the outside
of the collar. This kind of lid allows for the smallest possible bubble
in the container, as well as for the smallest amount of liquid lost to
In this photo the green liquid is the calibration buffer of pH=7.00. The probe's cap is also shown but not visible is a small moist sponge deep down.
Upon completion of each test, the vials are soaked and rinsed in clean
tap water or sea water. Note that sea water is less effective as it is
less solvent than fresh water, but it has no chlorine bleach which could
affect your results. It would be preferable to sterilise the canisters
with ultraviolet light, but letting them drip-dry upside down, then placed
on tissue paper, drying in bright sunlight achieves enough. Remember that
a small amount of contamination appears to be necessary to achieve complete
Do not use soap or other detergents, or bleach or any chemical, as this may upset the next test(s).
Using other containers? We have tested many types of container, small and large and came across some inexplicable differences. For some reason the plankton ecosystem is easily inhibited by PVC containers and even glass! So don't be dishearted if you don't get any results with different containers. First work with the FUJI HDPE film can before trying others.
A long measuring tape is needed for research that relates biodensity and rate of attack to underwater visibility. The idea behind it is that plant plankton must absorb light in order to photosynthesise. Thus it obscures light and is therefore visible. The more light is absorbed, the more chlorophyll is present, and there should be a close relationship. The theory behind this and practical results are explained in the chapter about the DDA method. Here we concern ourselves only with how to measure underwater visibility accurately.
In the scientific literature the Secchi disc method is described and used. This is a 20cm flat disc with 2 opposite quarters painted white and two black. It is lowered in a horizontal position off the measuring tape. Where the black and white pattern disappears, the depth is read from the tape. This method has several problems.
The most accurate measurements are done by floating in the sea in a wetsuit and observing the bag through a dark dive mask, for the following reasons:
When measuring inside estuaries where currents run, a long pole with a white object attached to its end is more suitable. See the chapter on equipment for suggestions.
The incubator is of critical importance to the DDA. One can modify a portable car refrigerator for this purpose as described in the chapter on equipment. Here we concern ourselves with how to use it. The travel incubator is a car refrigerator with the following alterations:
Keeping to a standardised time schedule is of critical importance because:
day of collection
Please note that improvements to the DDA technique are made from time to time and that this time schedule may change accordingly. Check this page from time to time.
Here is what a DDA card could look like on front and reverse side:
The DDA method has introduced the hion as a new unit for biodensity to facilitate comparisons. During decomposition, hydrogen ions are freed and many of these become hydrogen ions in solution, which are measured with a pH meter. The DDA method is based on the assumption that these hydrogen ions are a proxy for biomass although how the two relate EXACTLY, has not been ascertained. This chapter shows you how to calculate hion biodensity from the differences in pH.
The hion as unit of biodensity assumes that for most practical situations, the pH does not rise above 9.0, and if it does, the number of hions involved are so small (less than one) that they can be neglected, but fractions of a hion are still technically possible, even if they are not measurable with a three-digit pH meter.
The biodensity decomposed is calculated by subtracting the beginning value from the ending value, with proper ANTILOG (ALOG) conversion. It is not necessary to understand logarithms to do this correctly. Here is the formula where fpH= final pH and ipH= initial pH:
biodensity (hion) = ALOG( - fpH) - ALOG ( -ipH) in parts per billion (10^-9)So if the biodensity on the calculator reads 2.34 x 10^-7 it means 23.4 x 10^-8 or 234 x 10^-9 or 234 hion
On your calculator, use the memory function to store the initial value
and press the following buttons:
|type this||press this||what happens|
|8.06||+/-||the initial pH is negated to -8.06 which is what pH really stands for|
|ALOG||you may need to press two buttons like SHIFT and LOG to achieve this: 8.7096^-09|
|Min||store into memory. Some calculators use STO ore something.|
|6.88||+/-||enter final pH and negate it: -6.88|
|ALOG||anti-log it so the calculator reads: 1.3183^-07|
|- Mr =||subtract what is stored in memory and press the = button for the result:
which means 123^-09 or 123 hion
Before using the vials for testing, and after completion of each test, vials must be washed and dried. It is really very simple. Ideally one would want to sterilise them but this is not easily done without introducing new problems. Remember that each sample will contain a living ecosystem with thousands of species and millions of individuals. Remember also that the plankton ecosystem is easily inhibited by chemicals such as fluoride and chlorine from tap water and various kinds of soap.
If you are able to collect rain water from your roof, this is a real asset. If you have access to distilled water, it comes in handy. But you can achieve satisfactory results by simply rinsing the vials in warm tap water. Don't get tempted to use soap but wash your hands with soap first because a smudge of skin grease on the inside of a vial can upset measurements.
The rule for washing or rinsing is successive dilution. If your
first wash is effective only by 90%, leaving 10% 'dirt', the next wash
will leave 10% of 10% or 1%, and the next one after that 0.1% 'dirt'. So
the secret is to wash at least twice.
The third wash can be done with distilled water but this is not strictly necessary. It is more important to dry the containers completely such that pollutants like chlorine and fluoride evaporate. Shake the moisture off and place containers on a towel in the sunlight before a window. This provides enough ultraviolet light to reach the cleanliness desired for the next test. Once the containers and their caps are dry, cap them and store them away in the dark.
The ethyl alcohol solution used is diluted to a standard strength of 20% for the following reasons:
From the local chemist you can purchase 90% pure ethyl alcohol (normal
alcohol) which needs to be diluted 4.5 times for 20% final strength. You
can do this with either a sight glass or with scales. You won't need more
than 50ml of it.
Fill the sight glass to 10ml with 90% alchohol, then add distilled water to 45ml (35ml added). With nearly 30 drops per ml, this suffices for 1350 drops or at most 450 samples. Make a new solution when less than 20% or 10ml remain in the dispenser.