US Long-Term Ecological Research Network

LTREB Biological Limnology at Lake Myvatn 2012-current

Abstract
These data are part of a long-term monitoring program in the central part of Myvatn that represents the dominant habitat, with benthos consisting of diatomaceous ooze. The program was designed to characterize import benthis and pelagic variables across years as midge populations varied in abundance. Starting in 2012 samples were taken at roughly weekly inervals during June, July, and August, which corresponds to the summer generation of the dominant midge,<em>Tanytarsus gracilentus</em>.
Creator
Dataset ID
296
Date Range
-
Maintenance
Ongoing
Metadata Provider
Methods
Benthic Chlorophyll Field sampling (5 samples) (2012, 2013)1. Take 5 cores from the lake2. Cut the first 0.75 cm (1 chip) of the core with the extruder and place in deli container. Label with date and core number.3. Place deli containers into opaque container (cooler) and return to lab. This is the same sample that is used for the organic matter analysis.In 2014, the method for sampling benthic chlorophyll changed. The calculation of chlorophyll was changed to reflect the different area sampled. Below is the pertinent section from the methods protocols. Processing after the collection of the sample was not changed.Take sediment samples from the 5 cores collected for sediment characteristics. Take 4 syringes of sediment with 10mL syringe (15.96mm diameter). Take 4-5cm of sediment. Then, remove bottom 2cm and place top 2cm in the film canister.Filtering1. Measure volume of material in deli container with 60mL syringe and record.2. Homogenize and take 1mL sample with micropipette. The tip on the micropipette should be cut to avoid clogging with diatoms. Place the 1mL sample in a labeled film canister. Freeze sample at negative 20 degrees Celsius unless starting methanol extraction immediately.3. Add 20mL methanol. This methanol can be kept cool in the fridge, although then you will need a second bottle of methanol for the fluorometer. Shake for 5 sec.4. After 6-18 hours, shake container for 5 sec.Fluorometer1. Allow the film canisters to sit at room temperature for approximately 15 min to avoid excessive condensation on the glass tubes. Shake tubes for 5 sec after removing from fridge but then be careful to let them settle before removing sample.2. Record the sample information for all of the film canisters on the data sheet.3. Add 4mL of sample to a 13x100mL glass tube.4. Insert the sample into the fluorometer and record the reading in the Fluor Before Acid column. The sample reading should be close to one of the secondary solid standards (42ug/L or 230ug/L), if not, dilute the sample to within 25 per cent of the secondary solid standards (30-54ug/L or 180-280ug/L). It is a good idea to quickly check 2mL of a sample that is suspected to be too high to get an idea if other samples may need to be diluted. If possible, read the samples undiluted.5. If a sample needs to be diluted, use a 1000 microLiter pipette and add 2mL of methanol to a tube followed by 2mL of undiluted sample. Gently invert the tube twice and clean the bottom with a paper towel before inserting it into the fluorometer. If the sample is still outside of the ranges above, combine 1 mL of undiluted sample with 3 mL of methanol. Be sure to record the dilution information on the data sheet.6. Acidify the sample by adding 120microLiters of 0.1 N HCl (30microLiters for every one mL of sample). Then gently invert the sample and wait 90 seconds (we used 60 seconds in 2012, the protocol said 90) before putting the sample into the fluorometer and recording the reading in the Fluor After Acid column. Be sure to have acid in each tube for exactly the same amount of time. This means doing one tube at a time or spacing them 30-60 seconds apart.7. Double check the results and redo samples, which have suspicious numbers. Make sure that the after-acidification values make sense when compared to the before acidification value (the before acid/after acid ratio should be approximately the same for all samples).Clean up1. Methanol can be disposed of down the drain as long as at least 50 times as much water is flushed.2. Rinse the film canisters and lids well with tap water and scrub them out with a bottle brush making sure to remove any remaining filter paper. Give a final rinse with distilled water. Pelagic Chlorophyll Field sampling (5 samples)1. Take 2 samples at each of three depths, 1, 2, and 3m with Arni&rsquo;s zooplankton trap. For the 1m sample, drop the trap to the top of the chain. Each trap contains about 2.5L of water when full. 2. Empty into bucket by opening the bottom flap with your hand.3. Take bucket to lab.Filtering1. Filter 1L water from integrated water sample (or until the filter is clogged) through the 47 mm GF/F filter. The pressure used during filtering should be low ( less than 5 mm Hg) to prevent cell breakage. Filtering and handling of filters should be performed under dimmed lighting.2. Remove the filter with forceps, fold it in half (pigment side in), and put it in the film canister. Take care to not touch the pigments with the forceps.3. Add 20mL methanol. This methanol can be kept cool in the fridge, although then you will need a second bottle of methanol for the fluorometer. Shake for 5 sec. and place in fridge.4. After 6-18 hours, shake container for 5 sec.5. Analyze sample in fluorometer after 24 hours.Fluorometer1. Allow the film canisters to sit at room temperature for approximately 15 min to avoid excessive condensation on the glass tubes. Shake tubes for 5 sec after removing from fridge but then be careful to let them settle before removing sample.2. Record the sample information for all of the film canisters on the data sheet.3. Add 4mL of sample to a 13x100mL glass tube.4. Insert the sample into the fluorometer and record the reading in the Fluor Before Acid column. The sample reading should be close to one of the secondary solid standards (42ug/L or 230ug/L), if not, dilute the sample to within 25 percent of the secondary solid standards (30-54ug/L or 180-280ug/L). It is a good idea to quickly check 2mL of a sample that is suspected to be too high to get an idea if other samples may need to be diluted. If possible, read the samples undiluted.5. If a sample needs to be diluted, use a 1000uL pipette and add 2mL of methanol to a tube followed by 2mL of undiluted sample. Gently invert the tube twice and clean the bottom with a paper towel before inserting it into the fluorometer. If the sample is still outside of the ranges above, combine 1 mL of undiluted sample with 3 mL of methanol. Be sure to record the dilution information on the data sheet.6. Acidify the sample by adding 120 microLiters of 0.1 N HCl (30 microLiters for every one mL of sample). Then gently invert the sample and wait 90 seconds (we used 60 seconds in 2012, the protocol said 90) before putting the sample into the fluorometer and recording the reading in the Fluor After Acid column. Be sure to have acid in each tube for exactly the same amount of time. This means doing one tube at a time or spacing them 30-60 seconds apart.7. Double check the results and redo samples, which have suspicious numbers. Make sure that the after-acidification values make sense when compared to the before acidification value (the before acid/after acid ratio should be approximately the same for all samples).Clean up1. Methanol can be disposed of down the drain as long as at least 50 times as much water is flushed.2. Rinse the film canisters and lids well with tap water and scrub them out with a bottle brush making sure to remove any remaining filter paper. Give a final rinse with distilled water. Pelagic Zooplankton Counts Field samplingUse Arni&rsquo;s zooplankton trap (modified Schindler) to take 2 samples at each of 1, 2, and 3m (6 total). For the 1m sample, drop the trap to the top of the chain. Each trap contains about 2.5L of water when full. Integrate samples in bucket and bring back to lab for further processing.Sample preparation in lab1. Sieve integrated plankton tows through 63&micro;m mesh and record volume of full sample2. Collect in Nalgene bottles and make total volume to 50mL3. Add 8 drops of lugol to fix zooplankton.4. Label bottle with sample date, benthic or pelagic zooplankton, and total volume sieved. Samples can be stored in the fridge until time of countingCounting1. Remove sample from fridge2. Sieve sample with 63 micro meter mesh over lab sink to remove Lugol&rsquo;s solution (which vaporizes under light)3. Suspend sample in water in sieve and flush from the back with squirt bottle into counting tray4. Homogenize sample with forceps or plastic pipette with tip cut off5. Identify (see zooplankton identification guide) using backlit microscope and count with multiple-tally counter. i. Set magnification so that you can see both top and bottom walls of the tray. ii. Change focus depth to check for floating zooplankton that must be counted as well.6. Pipette sample back into Nalgene bottle, add water to 50mL, add 8 drops Lugol&rsquo;s solution, and return to fridgeSubsamplingIf homogenized original sample contains more than 500 individuals in the first line of counting tray, you may subsample under the following procedure.1. Return original sample to Nalgene bottle and add water to 50mL2. Homogenize sample by swirling Nalgene bottle3. Collect 10mL of zooplankton sample with Hensen-Stempel pipette4. Empty contents of Hensen-Stempel pipette into large Bogorov tray5. Homogenize sample in tray with forceps or plastic pipette with tip cut off6. Identify (see zooplankton identification guide) using backlit microscope and count with multiple-tally counter. i. Set magnification so that you can see both top and bottom walls of the tray. ii. Change focus depth to check for floating zooplankton that must be counted, too! 7. Pipette sample back into Nalgene bottle, add water to 50mL, add 8 drops Lugol&rsquo;s solution, and return to fridge Benthic Microcrustacean Counts Field samplingLeave benthic zooplankton sampler for 24h. Benthic sampler consists of 10 inverted jars with funnel traps in metal grid with 4 feet. Set up on bench using feet (on side) to get a uniform height of the collection jars (lip of jar = 5cm above frame). Upon collection, pull sampler STRAIGHT up, remove jars, homogenize in bucket and bring back to lab. Move the boat slightly to avoid placing sampler directly over cored sediment.Sample preparation in lab1. Sieve integrated samples through 63 micrometer mesh and record volume of full sample2. Collect in Nalgene bottles and make total volume to 50mL3. Add 8 drops of lugol to fix zooplankton.4. Label bottle with sample date, benthic or pelagic zooplankton, and total volume sieved. Samples can be stored in the fridge until time of countingCounting1. Remove sample from fridge2. Sieve sample with 63 micrometer mesh over lab sink to remove Lugol&rsquo;s solution (which vaporizes under light)3. Suspend sample in water in sieve and flush from the back with squirt bottle into counting tray4. Homogenize sample with forceps or plastic pipette with tip cut off5. Identify (see zooplankton identification guide) using backlit microscope and count with multiple-tally counter. i. Set magnification so that you can see both top and bottom walls of the tray. ii. Change focus depth to check for floating zooplankton that must be counted, too!6. Pipette sample back into Nalgene bottle, add water to 50mL, add 8 drops Lugol&rsquo;s solution, and return to fridgeSubsamplingIf homogenized original sample contains more than 500 individuals in the first line of counting tray, you may subsample under the following procedure.1. Return original sample to Nalgene bottle and add water to 50mL2. Homogenize sample by swirling Nalgene bottle3. Collect 10mL of zooplankton sample with Hensen-Stempel pipette4. Empty contents of Hensen-Stempel pipette into large Bogorov tray5. Homogenize sample in tray with forceps or plastic pipette with tip cut off6. Identify (see zooplankton identification guide) using backlit microscope and count with multiple-tally counter. i. Set magnification so that you can see both top and bottom walls of the tray. ii. Change focus depth to check for floating zooplankton that must be counted, too! 7. Pipette sample back into Nalgene bottle, add water to 50mL, add 8 drops Lugol&rsquo;s solution, and return to fridge Chironomid Counts (2012, 2013) For first instar chironomids in top 1.5cm of sediment only (5 samples)1. Use sink hose to sieve sediment through 63 micrometer mesh. You may use moderate pressure to break up tubes.2. Back flush sieve contents into small deli container.3. Return label to deli cup (sticking to underside of lid works well).For later instar chironomids in the section 1.5-11.5cm (5 samples)4. Sieve with 125 micrometer mesh in the field.5. Sieve through 125micrometer mesh again in lab to reduce volume of sample.6. Transfer sample to deli container or pitfall counting tray.For all chironomid samples7. Under dissecting scope, pick through sieved contents for midge larvae. You may have to open tubes with forceps in order to check for larvae inside.8. Remove larvae with forceps while counting, and place into a vial containing 70 percent ethanol. Larvae will eventually be sorted into taxonomic groups (see key). You may sort them into taxonomic groups as you pick the larvae, or you can identify the larvae while measuring head capsules if chironomid densities are low (under 50 individuals per taxanomic group).9. For a random sample of up to 50 individuals of each taxonomic group, measure head capsule, see Chironomid size (head capsule width).10. Archive samples from each sampling date together in a single 20mL glass vial with screw cap in 70 percent ethanol and label with sample contents , Chir, sample date, lake ID, station ID, and number of cores. Chironomid Cound (2014) In 2014, the method for sampling chironomid larvae changed starting with the sample on 2014-06-27; the variable &quot;top_bottom&quot; is coded as a 2. In contrast to previous measurements, the top and bottom core samples were combined and then subsampled. Below is the pertinent section of the protocols.Chironomid samples should be counted within 24 hours of collection. This ensures that larvae are as active and easily identified as possible, and also prevents predatory chironomids from consuming other larvae. Samples should be refrigerated upon returning from the field.<strong>For first instar chironomids in top 1.5cm of sediment only (5 samples)</strong>1. Use sink hose to sieve sediment through 63&micro;m mesh. You may use moderate pressure to break up tubes.2. Back flush sieve contents using a water bottle into small deli container.3. Return label to deli cup (sticking to underside of lid works well).<strong>For larger instar chironomids in the section 1.5-11.5cm (5 samples)</strong>4. Sieve with 125&micro;m mesh in the field.5. Sieve through 125&micro;m mesh again in lab to reduce volume of sample and break up tubes.6. Transfer sample to deli container with the appropriate label.<strong>Subsample if necessary</strong>If necessary, subsample with the following protocol.a. Combine top and bottom samples from each core (1-5) in midge sample splitter.b. Homogenize sample thoroughly, collect one half in deli container, and label container with core number and &ldquo;1/2&rdquo;c. If necessary, split the half that remains in the sampler into quarters, and collect each in deli containers labeled with core number, &ldquo;1/4&rdquo;, and replicate 1 or 2d. Store all deli containers in fridge until counted, and save until all counting is complete&quot; Chironomid Size (head capsule width) 1. Obtain picked samples preserved in ethanol and empty onto petri dish.2. Sort larvae by family groups, arranging in same orientation for easy measurment.3. Set magnification to 20, diopter, x 50 times4. Take measurments for up to 50 or more individuals of each taxa. Round to nearest optical micrometer unit.5. Fill out data sheet for number of larvae in each taxa, Chironomid measurements for each taxa, date of sample, station sample was taken from, which core the sample came from, who picked the core, and your name as the measurer.6. Enter data into shared sheetSee &quot;Chironomid Counts&quot; for changes in sampling chironomid larvae in 2014.
Version Number
17

North Temperate Lakes LTER: Chlorophyll - Trout Lake Area 1981 - current

Abstract
Chlorophyll and phaeopigments are measured at our permanent sampling station in the deepest part of each lake. Chlorophyll samples are collected from the seven primary study lakes (Allequash, Big Muskellunge, Crystal, Sparkling, and Trout lakes and bog lakes 27-02 [Crystal Bog], and 12-15 [Trout Bog]) in the Trout Lake area at two to 10 depths depending on the lake and analyzed spectrophotometrically. Sampling Frequency: fortnightly during ice-free season - every 6 weeks during ice-covered season Number of sites: 7
Core Areas
Dataset ID
35
Date Range
-
LTER Keywords
Maintenance
ongoing
Metadata Provider
Methods
Spectrophotometer:A. Chlorophyll Extraction (using tissue grinder at DNR Research Station) 1. Dim the lights and keep the sample tubes in the freezer: Because chlorophyll degrades when exposed to light and heat, this procedure and all others associated with analyzing chlorophyll should be carried out in dim light conditions. Only one sample tube should be out of the freezer at any one time while the pre-grinding or grinding procedure is occurring. Return each tube to the freezer as soon as its filter has been ground. 2. Pre-grind filters: Use the sharpened stainless steel probe to chop up the filter into small pieces. This should take approximately 2 minutes. 3. Grind filters: The teflon tip on the tissue grinder should be sanded after grinding approximately 5 filters. Grind each filter for 2 minutes. Do not lift the teflon tip out of the test tube while the grinder is rotating. Grind the filters by attempting to keep the teflon tip in the acetone solution and pressing the tip against the filter and the tube. 4. Return the sample tubes to the freezer for 24 hours: Most protocols call for extracting the samples in the refrigerator (at 4 degrees C). However, after extracting duplicate samples in the freezer and refrigerator (after grinding) there was no significant difference in the chlorophyll results. Because past samples have been extracted in the freezer, this is the current procedure being used. B. Centrifuging the Samples: The samples should be centrifuged as close as possible to 24 hours after extraction. Before centrifuging the samples, turn on the spectrophotometer and enter the correct program number to be sure that it is working properly. Perform the procedures below in dim light. 1. Checking acetone volume: In dim light, use an identical tube as those used for the samples but with mL marked on it, to measure the volume of the acetone in the samples. Measure to the nearest 0.5 mL. If the sample has any other volume than 5 mL, write the volume on the sample label and remember to enter the volume later into the spreadsheet. 2. Loading the centrifuge: Making sure that the rubber stoppers are on tight, put tubes with equal acetone volumes opposite each other in the centrifuge. If there is an odd tube remaining or a tube with a different volume, put a spare tube opposite the sample with the same volume of water to counterbalance the centrifuge. 4. Running the centrifuge: Turn the speed dial below 40. Turn the timer past 15 minutes. Slowly turn up the speed allowing time for the centrifuge to increase in speed. If there is an imbalance in the centrifuge (or any other problem), the centrifuge will run much louder than normal. In this case, stop the centrifuge and attempt to locate the imbalance. If the centrifuge is running smoothly, set the speed at 90 and the timer at 15 minutes. Previously, the numbers on the dial were believed to correspond to revolutions per second; however, this is not the case, for the centrifuge will only reach rpms of approximately 2500. 5. Unloading the centrifuge: Allow the centrifuge to come to a stop on its own. Carefully take each sample tube out of the centrifuge with minimal mixing. If the filter paper is mixed with the liquid, it will be necessary to re-centrifuge the sample. Transport the samples to the spectrophotometer in a rack that has tinfoil on the sides in order to block out the light. C. Running a Sample: 1. Select the test: Allow the spectrophotometer to warm up for at least 15 minutes. Select the proper program by pressing the test number followed by Select. 2. Rinse the cuvettes 3 times with acetone. It is most efficient to rotate 4 matching 1 cm cuvettes. Try to touch the cuvettes only on the opaque sides avoiding touching the clear sides especially on the lower half of the cuvette. 3. Run a blank and check that all cuvettes read near 0: Add acetone to the 4 matching cuvettes (at least half full), wipe them clean with a tissue, and insert them into the spectrophotometer with the labeled sides all facing the same direction (always put the tops on the cuvettes when they are in the spec). Press Run and the spec. will ask for a blank. Use one of the cuvettes filled with acetone as the blank. Once the blank is run, run all of the cuvettes (the cuvette position is changed by pulling out the metal rod to the next notched position). All of the readings at all wavelengths should be within .001 of 0. If this is not the case, remove the suspect cuvette and rinse, wipe, add acetone, and rerun it. Make sure that the correct program is being run by checking the wavelengths. The LTER samples should be run at 750, 665, 664, 647, and 630 nm. 4. Rinse the pipette tip: Before adding sample to a cuvette, the pipette tip should be rinsed with acetone. You should have 2 different sized beakers, one for waste and one for acetone rinse. Set the 10-1000uL pipette to 1000 uL (1 mL) and pipette 1mL of acetone from the rinse beaker and dispose of it in the waste beaker. Be sure that the pipette tip is firmly on the pipette (press it on the bottom of the rinse beaker). 5. Add sample to a cuvette: Before bringing the samples into the spectrophotometer room, turn off the overhead light and turn on the desk light in the corner. Carefully remove a sample from the rack and pipette approximately 2 mL of sample into a cuvette. Use caution not to suck up any filter paper into the pipette; tilt the sample to the side and submerge the pipette tip only just below the fluid level. If the pipette tip is getting close to the filter paper when removing the second mL of sample, stop pipetting and add the partial mL to the cuvette (it is possible to read approximately 1.5 mL of sample). 6. Check the 750 nm reading and run the sample: Insert the cuvette into the spec. (making sure that the labeled side is always facing in the same direction). The default reading on the spec is 750 nm. Check to make sure that this reading is less than 0.010 A. If the reading is higher, remove the cuvette and re-wipe it with a tissue. If the reading is still high, pour the sample back into the tube and re-centrifuge it. To run the sample press Run. 7. Acidify the sample: Once the sample has been run, remove it from the spec and add 60 uL of 0.1 N HCl (30 uL per 1 mL of sample). Gently shake the sample and wait 90 seconds to run it. 8. Check the acidification ratio: The before acidorafter acid ratio of the LTER samples is usually between 1.3 and 1.7. Compare the two readings to make sure the ratio fits in this range. If the ratio is higher than 1.7, re-acidify the sample and run it again (the acid probably did not make contact with the sample). 9. Rinse the cuvette: After checking the acidification ratio, dispose of the sample in the waste beaker and rinse the cuvette 3 times with acetone. Be sure to fill the cuvette to the top with acetone during each rinse to be sure that there is not any trace of acid left. Running Multiple Samples: 1. It may be more efficient to run 2 samples before acidification and then run them both after acidification. If this is done, take caution to add the correct sample to the correct cuvette and not to mix up the samples when they are removed from the spec. for acidification. Recording the Results: 1. Write the spec. id number located on the left of the printout onto the label of the corresponding sample. Each sample should have a before and an after acidification spec. id number written on its label. After all of the samples have been run, enter the date of analysis onto the spec. printout. This date will be used to identify the spec. printout when the data is proofread (after which proofed from spec. printout should be written on the spreadsheet). Clean-up: 1. Rinse the cuvettes 3 times with acetone, allow them to dry for several minutes in the cuvette rack, and return them to their box. 2. Solutions of less than 20percent Acetone can be disposed of down the drain followed by at least 10 volumes of water. Fill the waste beaker with water and pour the waste down the sink with the water running. Leave the water running for several minutes 3. Rinse the beakers and pipette tips 3 times with tap water followed by 3 rinses with distilled water. Hang the beakers on the drying rack. &nbsp;
Short Name
NTLPL01
Version Number
30

Little Rock Lake Experiment at North Temperate Lakes LTER: Physical Limnology 1983 - 2000

Abstract
The Little Rock Acidification Experiment was a joint project involving the USEPA (Duluth Lab), University of Minnesota-Twin Cities, University of Wisconsin-Superior, University of Wisconsin-Madison, and the Wisconsin Department of Natural Resources. Little Rock Lake is a bi-lobed lake in Vilas County, Wisconsin, USA. In 1983 the lake was divided in half by an impermeable curtain and from 1984-1989 the northern basin of the lake was acidified with sulfuric acid in three two-year stages. The target pHs for 1984-5, 1986-7, and 1988-9 were 5.7, 5.2, and 4.7, respectively. Starting in 1990 the lake was allowed to recover naturally with the curtain still in place. Data were collected through 2000. The main objective was to understand the population, community, and ecosystem responses to whole-lake acidification. Funding for this project was provided by the USEPA and NSF. Parameters characterizing the physical limnology of the treatment (north basin, stations 1 and 3) and reference basin (south basin, station 2 and 4) are usually measured at one station in the deepest part of each basin (stations 1 and 2) at 0.5 to 1-m depth intervals depending on the parameter. Parameters measured at depth include water temperature, vertical penetration of photosynthetically active radiation (PAR), dissolved oxygen, chlorophyll and phaeopigments. Additional derived parameters include fraction of surface PAR at each depth and percent oxygen saturation. Auxiliary data include time of day, air temperature, cloud cover, and wind speed and direction and secchi depth. Sampling Frequency: varies - Number of sites: 4
Core Areas
Dataset ID
248
Date Range
-
Maintenance
completed
Metadata Provider
Methods
Reading Temperature and Dissolved Oxygen1. Before leaving to sample a lake, check to make sure that there are no air bubbles under the probe membrane of the YSI TemperatureorDissolved Oxygen meter. If there are air bubbles or if it has been several months since changing the membrane (or if the instrument does not calibrate well or the oxygen readings wander), change the membrane as explained in the manual. Note: We have always used the Standard membranes. If adding water to new membrane fluid bottle (KCl), make sure to add Milli-Q water and not the CFL distilled water.2. Be sure to always store the probe in 100percent humidity surrounded by a wet sponge or paper towel.3. Turn on the temperatureordissolved oxygen meter at least 30 minutes before using it. It is best to turn it on before leaving to sample a lake as it uses up batteries slowly.4. Calibrate the meter using the chart on the back of the instrument (adjusted to the Madison altitude - 97percent oxygen saturation). Leave the plastic cap on the probe (at 100percent humidity). The temperature should not be changing during the calibration. Zero the instrument. When the temperature equilibrates, adjust the oxygen to correspond to the chart. After calibrating the instrument, switch the knob to percent oxygen saturation to make sure it is close to 97percent.5. Take readings at 1 meter intervals making sure to gently jiggle the cord when taking the oxygen readings (to avoid oxygen depletion). Jiggling the cord is not necessary if using a cable with a stirrer. Take half meter readings in the metalimnion (when temperature andoror oxygen readings exhibit a greater change with depth). A change of temperature greater than 1degreeC warrants half-meter intervals.6. Record the bottom depth using the markings on the temp.oroxygen meter cord and take a temperature and dissolved oxygen reading with the probe lying on the lake bottom. Dont forget to jiggle the probe to remove any sediment.7. If any readings seem suspicious, check them again when bringing the probe back up to the surface. You can also double check the calibration after bringing the probe out of the water (and putting the cap back on). Light (PAR) extinction coefficient is calculated by linearly regressing ln (FRLIGHT (z)) on depth z where the intercept is not constrained. FRLIGHT(z) = LIGHT(z) or DECK(z) where LIGHT(z) is light measured at depth z and DECK(z) is light measured on deck (above water) at the same time.For open water light profiles, the surface light measurement (depth z = 0) is excluded from the regression.For winter light profiles taken beneath the ice, the first light data are taken at the bottom of the ice cover and are included in the regression. The depth of uppermost light value is equal to the depth of the ice adjusted by the water level in the sample hole, i.e., the depth below the surface of the water. The water level can be at, above or below the surface of the ice. If the water level was not recorded, it is assumed to be 0.0 and the calculated light extinction coefficient is flagged. If ice thickness was not recorded, a light extinction coefficient is not calculated.For light data collected prior to March, 2007, light values less than 3.0 (micromolesPerMeterSquaredPerSec) are excluded. For light data collected starting in March 2007, light values less than 1.0 (micromolesPerMeterSquaredPerSec) are excluded. Except for bog lakes before August 1989, a light extinction coefficient is not calculated if there are less than three FRLIGHT values to be regressed. For bog lakes before August 1989, a light extinction coefficient is calculated if there are least two FRLIGHT values to be regressed. In these cases, the light extinction coefficient is flagged as non-standard.FRLIGHT values should be monotonically decreasing with depth. For light profiles where this is not true, a light extinction coefficient is not calculated.For samples for which light data at depth are present, but the corresponding deck light are missing, a light extinction coefficient is calculated by regressing ln (LIGHT (z)) on depth z. Note that if actual deck light had remained constant during the recording of the light profile, the resulting light extinction coefficient is the same as from regressing ln(FRLIGHT(z)). In these cases, the light extinction coefficient is flagged as non-standard.
Short Name
LRPHYS1
Version Number
4

Landscape Position Project at North Temperate Lakes LTER: Chlorophyll 1998 - 2000

Abstract
Parameters characterizing the chemical limnology and spatial attributes of 49 lakes were surveyed as part of the Landscape Position Project. Most parameters are measured at or close to the deepest part of the lake. Chlorophyll is measured by collecting separate integrated samples from the epilimnion, metalimnion, and hypolimnion Sampling Frequency: generally monthly for one summer; for some lakes, one or two samples in one summer Number of sites: 51
Core Areas
Dataset ID
92
Date Range
-
LTER Keywords
Maintenance
completed
Metadata Provider
Methods
Chlorophyll is measured by collecting separate integrated samples from the epilimnion, metalimnion, and hypolimnion
Short Name
LPPCHL1
Version Number
8

Cascade Project at North Temperate Lakes LTER: Process Data 1984 - 2007

Abstract
Data on chlorophyll, primary productivity, and alkaline phosphatase activity from 1984-2007. Samples were collected with a Van Dorn bottle at 6 depths determined from the percent of surface irradiance (100%, 50%, 25%, 10%, 5% and 1%) and in the hypolimnion (12 m in Peter, East Long, West Long, and Tuesday lakes; 9 m in Paul Lake; and 4.5 m in Central Long Lake). Sampling Frequency: varies Number of sites: 8
Core Areas
Dataset ID
73
Date Range
-
LTER Keywords
Maintenance
completed
Metadata Provider
Methods
CHLOROPHYLL a ANALYSISEQUIPMENT: Film canistersTurner 450 Fluorometer fitted with:1. Quartz-halogen lamp2. Emission filter -SC6653. Excitation filter -NB44047mm Whatman GForF filters12 x 75 mm disposable glass culture cuvettes (Do not reuse cuvettes!)1-5 mL Oxford pipettorFinnpipette Stepper Pipetter with 5 mL tiptimestimesNOTEtimestimes-Change filters with fluorometer off! (Remember that chlorophyll analysis filters are different from APA analysis filters.)-Make sure Fluorometer has been calibrated for chlorophyll a (see Fluorometer Calibration for Chlorophyll a Analysis).REAGENTS: 100percent Methanol, spectrophotometric gradeCAUTION - wear gloves whenever you use methanol.0.1 N HCLEthidium Bromide Stock 3 standard (40microM solution)PROCEDURE:A. Filter water samples from each of the 6 light-depths onto a 47 mm GForF filter.1. Filters have a grid side and a smooth side. Place filter smooth side up.2. Shake sample bottle well before filtering (do this after the DIC sample has been taken from the same bottle.)3. For each depth, filter enough water so there is a faint color on the filter. For our lakes this ranges between 100-300ml. Record the volume filtered. Make sure you filer at less than 200 mm Hg pressure.4. Rinse filter towers and filters with DI water, place filters in labeled film canisters and place in freezer. Labels should include lake, date, and depth ID.5. If measuring edible chlorophyll as well, repeat steps 1-4 above, but first filter the sample through 35 microm mesh. (This has not been done since 2001, inclusive.)B. Extraction - DO IN DIM LIGHT and WEAR GLOVES!!1. Remove one tray of film canisters from the freezer. Extract chlorophyll by adding 25 mL 100percent MeOH to each film canister. If using re-pipettor, verify dispensed volume. (Record extraction volume if different from 25 mL.) Note the extraction time for each group of samples.2. Re-cap and place canisters in refrigerator to extract for exactly 24 hours (in the dark).3 Repeat steps 1 and 2 for all trays that have been in the freezer more than 24 hours.C. FluorometryCalibration of the fluorometer using a chlorophyll standard is typically performed at the beginning of the field season, or when a bulb is changed. Calibration using Ethidium Bromide is done at the beginning of each sample set.1. Insert correct filters in fluorometer while fluorometer is off. (Emission filter -SC665, Excitation filter -NB440), and warm it up for 1 hour .2. TURN LIGHTS OUT. Chlorophylls must be read in low light and samples must be kept cool. Do not remove film canisters from the refrigerator until you are ready to process the samples.3. Place clean cuvettes into a labeled rack (12 cuvettes per rack). Remove one lake-day of film canisters from the refrigerator.4. Place Ethidium Bromide Stock 3 standard into fluorometer and record reading on datasheet. Then, turn the span knob until the reading is 908. Record this on the datasheet.5. Shake film canister, remove the lid, and rinse the pipette tip with 2.5 mL of the sample. Then remove 2.5 mL of sample and place in cuvette.times Repeat for all film canisters.6. Pipette 2.5 mL of 100percent methanol into a cuvette for the blank and use it to zero the fluorometer. Choose a gain and turn the zero knob until the fluorometer reads 000. You must zero the machine every time you change gains.7. Remove the first sample cuvette from the rack, wipe with a Kimwipe, and place in fluorometer. Record the gain and the fluorescence before acidification, Fb. Repeat for all 12 cuvettes in the rack. Readings should be between about 200 and 1000. If not, adjust the gain and re-zero.8. Acidify each cuvette with 100 microL 0.0773 N HCl using the repeating pipetter and mix (hold the top of the cuvette securely, then &quot;thump&quot; the bottom several times). Check for condensation on the outside of the cuvettes, and wipe with a Kimwipe if necessary. Wait about 1 min from the acidification of the first cuvette.9. Record the fluorescence after acidification for all 12 cuvettes. VERY IMPORTANT: Make sure you read the Fb and Fa values for each sample on the same gain.10. Remove a new lake-day batch of film canisters from the refrigerator and repeat steps 3-9.times if particulate matter is present, centrifuge sample for 10 min. and use supernatant.D. Clean Up: DO THIS UNDER THE HOOD!1. Dump methanol solution from cuvettes and film canisters into a metal tray. Place the film canisters and lids in a separate tray. Position them in one layer on the tray with their openings facing up. Leave the trays under the hood overnight to evaporate the methanol.REFERENCES:Marker, A.F.H., C.A. Crowther, and R.J.M. Gunn. 1980. Methanol and acetone as solvents for estimating chlorophyll a and phaeopigments by spectrophotometry. Arch. Hydrobiol. Beih. Ergebn. Limnol 14: 52-69.Strickland, J.H. and T.R. Parsons. 1968. A practical handbook of seawater analysis. Fish. Res. Brd. Can. Bulletin 167.pp. 201-206.Holm-Hansen, O. 1978. Chlorophyll a determination: improvements in methodology. Oikos 30:438-447.
Short Name
CPROC1
Version Number
6

Lake Wingra Exclosure Experiment at North Temperate Lakes LTER: Chlorophyll 2005 - 2008

Abstract
Starting in late summer 2005, Wisconsin Dept of Natural Resources (WDNR), Dane County, Friends of Lake Wingra (FOLW), and NTL-LTER initiated a 3-year experiment in Lake Wingra to test the response of the native macrophyte community to clearer water produced from a major carp reduction program. This demonstration-scale experiment includes the construction of a 1.0-hectare rectangular carp exclosure with its solid vinyl walls extending from the lake shoreline to a water depth of 2.9 meters. NTL-LTER conducts the routine limnological monitoring of the lake and exclosure and is leading the science evaluation of potential lake restoration activities. The exclosure experiment was terminated in the fall of 2008. The exclosure was removed from Lake Wingra at that time. Sampling is done both within the exclosure and at a control site located nearby in the littoral zone. The sample location within the exclosure is equidistant from the side walls and approximately 75 meters from the shore in a water depth of approximately 2.5 meters. The control site sample location is approximately 75 meters west of the exclosure sample site at the same approximate distance from shore and water depth. Samples are taken at the same time and on the same schedule as the NTL-LTER limnological sampling on Lake Wingra, e.g., biweekly spring through summer, every 4 weeks in the fall, and once during the winter depending on ice conditions. Parameters measured within the exclosure and at the control site include water temperature, dissolved oxygen, secchi depth and chlorophyll-a. Additional parameters measured only within the exclosure include total Kjeldahl nitrogen, nitrate + nitrite nitrogen, ammonia nitrogen, total phosphorus, dissolved reactive phosphorus and dissolved reactive silica. Chlorophyll is measured within the exclosure and at a nearby control site in the littoral zone. Spectrophotometric analysis and fluorometric analysis are done on integrated samples from surface to 1 meter. The first data table below, Chlorophyll - Tri Chlor Spec, contains only the Tri_chlor_spec values for the exclosure and control samples. This measurement is the sum of chlorophyll a concentration and phaeophyton concentration using a spectrophotometer. The second data table, Chlorophyll - Full Series, contains all the results of the spectrophotometric analysis and fluorometric analysis. Sampling Frequency: generally bi-weekly during ice-free season from late March or early April through early September, then every 4 weeks through late November. Number of sites: 2
Core Areas
Dataset ID
190
Date Range
-
LTER Keywords
Maintenance
completed
Metadata Provider
Methods
Chlorophyll is measured within the exclosure and at a nearby control site in the littoral zone. Spectrophotometric analysis and fluorometric analysis are done on integrated samples from surface to 1 meter.
Short Name
FOLWEXCH
Version Number
22

North Temperate Lakes LTER: Chlorophyll - Madison Lakes Area 1995 - current

Abstract
Chlorophyll is measured at our permanent sampling station in the deepest part of the lake. Chlorophyll samples are collected from the four primary study lakes in the Madison area (Lakes Mendota, Monona, and Wingra and Fish Lake) at integrated depths and discrete depths for spectrophotometric analysis and fluorometric analysis. Due to a change in instruments starting in 2002 and lasting through 2007, chlorophyll analyses for the southern lakes had an uncorrectable bias, and are not included in this dataset. Analyses since then (2008 onward) have been determined to not have this bias. Sampling Frequency: bi-weekly during ice-free season from late March or early April through early September, then every 4 weeks through late November; sampling is conducted usually once during the winter (depending on ice conditions). Number of sites: 4
Core Areas
Dataset ID
38
Date Range
-
LTER Keywords
Maintenance
ongoing
Metadata Provider
Methods
SpectrophotometerA. Chlorophyll Extraction (using tissue grinder at DNR Research Station) 1. Dim the lights and keep the sample tubes in the freezer: Because chlorophyll degrades when exposed to light and heat, this procedure and all others associated with analyzing chlorophyll should be carried out in dim light conditions. Only one sample tube should be out of the freezer at any one time while the pre-grinding or grinding procedure is occurring. Return each tube to the freezer as soon as its filter has been ground. 2. Pre-grind filters: Use the sharpened stainless steel probe to chop up the filter into small pieces. This should take approximately 2 minutes. 3. Grind filters: The teflon tip on the tissue grinder should be sanded after grinding approximately 5 filters. Grind each filter for 2 minutes. Do not lift the teflon tip out of the test tube while the grinder is rotating. Grind the filters by attempting to keep the teflon tip in the acetone solution and pressing the tip against the filter and the tube. 4. Return the sample tubes to the freezer for 24 hours: Most protocols call for extracting the samples in the refrigerator (at 4 degrees C). However, after extracting duplicate samples in the freezer and refrigerator (after grinding) there was no significant difference in the chlorophyll results. Because past samples have been extracted in the freezer, this is the current procedure being used. B. Centrifuging the Samples: The samples should be centrifuged as close as possible to 24 hours after extraction. Before centrifuging the samples, turn on the spectrophotometer and enter the correct program number to be sure that it is working properly. Perform the procedures below in dim light. 1. Checking acetone volume: In dim light, use an identical tube as those used for the samples but with mL marked on it, to measure the volume of the acetone in the samples. Measure to the nearest 0.5 mL. If the sample has any other volume than 5 mL, write the volume on the sample label and remember to enter the volume later into the spreadsheet. 2. Loading the centrifuge: Making sure that the rubber stoppers are on tight, put tubes with equal acetone volumes opposite each other in the centrifuge. If there is an odd tube remaining or a tube with a different volume, put a spare tube opposite the sample with the same volume of water to counterbalance the centrifuge. 4. Running the centrifuge: Turn the speed dial below 40. Turn the timer past 15 minutes. Slowly turn up the speed allowing time for the centrifuge to increase in speed. If there is an imbalance in the centrifuge (or any other problem), the centrifuge will run much louder than normal. In this case, stop the centrifuge and attempt to locate the imbalance. If the centrifuge is running smoothly, set the speed at 90 and the timer at 15 minutes. Previously, the numbers on the dial were believed to correspond to revolutions per second; however, this is not the case, for the centrifuge will only reach rpms of approximately 2500. 5. Unloading the centrifuge: Allow the centrifuge to come to a stop on its own. Carefully take each sample tube out of the centrifuge with minimal mixing. If the filter paper is mixed with the liquid, it will be necessary to re-centrifuge the sample. Transport the samples to the spectrophotometer in a rack that has tinfoil on the sides in order to block out the light. C. Running a Sample: 1. Select the test: Allow the spectrophotometer to warm up for at least 15 minutes. Select the proper program by pressing the test number followed by Select. 2. Rinse the cuvettes 3 times with acetone. It is most efficient to rotate 4 matching 1 cm cuvettes. Try to touch the cuvettes only on the opaque sides avoiding touching the clear sides especially on the lower half of the cuvette. 3. Run a blank and check that all cuvettes read near 0: Add acetone to the 4 matching cuvettes (at least half full), wipe them clean with a tissue, and insert them into the spectrophotometer with the labeled sides all facing the same direction (always put the tops on the cuvettes when they are in the spec). Press Run and the spec. will ask for a blank. Use one of the cuvettes filled with acetone as the blank. Once the blank is run, run all of the cuvettes (the cuvette position is changed by pulling out the metal rod to the next notched position). All of the readings at all wavelengths should be within .001 of 0. If this is not the case, remove the suspect cuvette and rinse, wipe, add acetone, and rerun it. Make sure that the correct program is being run by checking the wavelengths. The LTER samples should be run at 750, 665, 664, 647, and 630 nm. 4. Rinse the pipette tip: Before adding sample to a cuvette, the pipette tip should be rinsed with acetone. You should have 2 different sized beakers, one for waste and one for acetone rinse. Set the 10-1000uL pipette to 1000 uL (1 mL) and pipette 1mL of acetone from the rinse beaker and dispose of it in the waste beaker. Be sure that the pipette tip is firmly on the pipette (press it on the bottom of the rinse beaker). 5. Add sample to a cuvette: Before bringing the samples into the spectrophotometer room, turn off the overhead light and turn on the desk light in the corner. Carefully remove a sample from the rack and pipette approximately 2 mL of sample into a cuvette. Use caution not to suck up any filter paper into the pipette; tilt the sample to the side and submerge the pipette tip only just below the fluid level. If the pipette tip is getting close to the filter paper when removing the second mL of sample, stop pipetting and add the partial mL to the cuvette (it is possible to read approximately 1.5 mL of sample). 6. Check the 750 nm reading and run the sample: Insert the cuvette into the spec. (making sure that the labeled side is always facing in the same direction). The default reading on the spec is 750 nm. Check to make sure that this reading is less than 0.010 A. If the reading is higher, remove the cuvette and re-wipe it with a tissue. If the reading is still high, pour the sample back into the tube and re-centrifuge it. To run the sample press Run. 7. Acidify the sample: Once the sample has been run, remove it from the spec and add 60 uL of 0.1 N HCl (30 uL per 1 mL of sample). Gently shake the sample and wait 90 seconds to run it. 8. Check the acidification ratio: The before acidorafter acid ratio of the LTER samples is usually between 1.3 and 1.7. Compare the two readings to make sure the ratio fits in this range. If the ratio is higher than 1.7, re-acidify the sample and run it again (the acid probably did not make contact with the sample). 9. Rinse the cuvette: After checking the acidification ratio, dispose of the sample in the waste beaker and rinse the cuvette 3 times with acetone. Be sure to fill the cuvette to the top with acetone during each rinse to be sure that there is not any trace of acid left. Running Multiple Samples: 1. It may be more efficient to run 2 samples before acidification and then run them both after acidification. If this is done, take caution to add the correct sample to the correct cuvette and not to mix up the samples when they are removed from the spec. for acidification. Recording the Results: 1. Write the spec. id number located on the left of the printout onto the label of the corresponding sample. Each sample should have a before and an after acidification spec. id number written on its label. After all of the samples have been run, enter the date of analysis onto the spec. printout. This date will be used to identify the spec. printout when the data is proofread (after which proofed from spec. printout should be written on the spreadsheet). Clean-up: 1. Rinse the cuvettes 3 times with acetone, allow them to dry for several minutes in the cuvette rack, and return them to their box. 2. Solutions of less than 20percent Acetone can be disposed of down the drain followed by at least 10 volumes of water. Fill the waste beaker with water and pour the waste down the sink with the water running. Leave the water running for several minutes 3. Rinse the beakers and pipette tips 3 times with tap water followed by 3 rinses with distilled water. Hang the beakers on the drying rack. &nbsp;
Short Name
NTLPL04
Version Number
27
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