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ARCTIC LTER STREAMS PROTOCOL:

KUPARUK RIVER, OKSRUKUYIK CREEK, AND NEW STREAM REACHES

TABLE OF CONTENTS

Field Sampling Methods

Sample processing and analyses at Toolik

Laboratory analyses at MBL and other home institutions

FIELD METHODS

Field study sites

  1. Kuparuk River
    1. Dripper sites
      1. The original P dripper site was at the 0.0 km site, located about 1.7 km upstream from the Dalton Highway crossing.
      2. From 1985 to 1995, the P dripper has been at the 0.59 km site, located about 1.1 km upstream of the Dalton Highway crossing. In 1996 the P-dripper site was moved downstream to 1.4km.
      3. Nitrogen dripping was done on the Kuparuk in 1986 and 1989. It was done just downstream to the Dalton Highway crossing for logistical reasons.
    2. Discharge has historically been measured in many different locations. Good sites include the cross section directly under the pipeline and the narrow, straight reach just upstream of the 0.59 km dripper.
    3. Traditional nutrient/water chemistry collection sites in the Kuparuk are:
      1. -0.800 km (control)
      2. -0.177 km (control)
      3. 0.347 km (control)
      4. 0.564 km (control)
      5. 0.740 km (fertilized)
      6. 0.822 km (fertilized)
      7. 1.025 km (fertilized)
      8. Hershey Creek (tributary)
      9. 1.390 km (fertilized/recovery)
      10. 1.500 km (fertilized)
      11. 1.845 km (fertilized)
      12. 2.065 km (fertilized)
      13. 3.730 km (fertilized)
      14. 4.250 km (fertilized)
    4. Traditional epilithic algae (chlorophyll a) collection sites in the Kuparuk are:
      1. -0.300 km (control)
      2. -0.100 km (control)
      3. 0.000 km (control)
      4. 0.347 km (control)
      5. 0.740 km (fertilized)
      6. 1.025 km (fertilized)
      7. 1.390 km (fertilized/recovery)
      8. 1.500 km (fertilized)
      9. 2.000 km (fertilized)
      10. 3.000 km (fertilized)
      11. 4.000 km (fertilized)
    5. Traditional insect collection sites in the Kuparuk are:
      1. -1.000 km (control)
      2. -0.500 km (control)
      3. -0.400 km (control)
      4. -0.100 km (control)
      5. 1.000 km (fertilized)
      6. 1.400 km (fertilized)
      7. 2.000 km (fertilized)
      8. 2.500 km (fertilized)
    6. Fish collection and weir sites
      1. Kuparuk River young-of-the-year Arctic grayling are most easily collected in broad, slow pools or reaches, such as those at:
        1. -0.100 km (control)
        2. 0.000 km (control)
        3. 1.800 km (fertilized)
        4. 2.400 km (fertilized)
        5. 2.700 km (fertilized)
        6. 3.500 km (fertilized)
      2. Adults are sampled by angling throughout both reaches; generally, the fertilized zone is sampled downstream of the Dalton Highway crossing to about 5 km, while the control zone is sampled well upstream of the dripper.
      3. Weirs in the Kuparuk are deployed in different locales in each year, as determined by Dr. Linda Deegan.
  2. Oksrukuyik Creek
    1. The dripper site at Oks Creek, for both the P dripper and the N dripper (1991-1996), is at the 0.0 km site, approximately 2.7 km upstream of the Dalton Highway crossing.
    2. The discharge site at Oks Creek is about 10 m downstream of the Dalton Highway crossing. It is marked by flagged rebar stakes on opposite banks.
    3. Traditional nutrient/water chemistry collection sites in Oks Creek are:
      1. -0.737 km (control)
      2. -0.327 km (control)
      3. Wolf Creek (tributary)
      4. -0.136 km (control)
      5. 0.227 km (fertilized)
      6. 0.482 km (fertilized)
      7. 1.056 km (fertilized)
      8. 1.370 km (fertilized)
      9. 1.700 km (fertilized)
      10. 2.496 km (fertilized)
    4. Traditional epilithic algae (chlorophyll a) collection sites in Oks Creek are identical to the nutrient sites in Part 3 (above), except no epilithic algae is collected in the tributary stream, Wolf Creek.
    5. Traditional insect collection sites in Oks Creek are:
      1. -0.737 km (control)
      2. -0.550 km (control)
      3. -0.327 km (control)
      4. -0.267 km (control)
      5. -0.136 km (control)
      6. 0.227 km (fertilized)
      7. 0.482 km (fertilized)
      8. 1.056 km (fertilized)
      9. 1.370 km (fertilized)
      10. 1.700 km (fertilized)
      11. 2.496 km (fertilized)
    6. Fish collection and weir sites
      1. Oks Creek young-of-the-year Arctic grayling are most easily collected in broad, slow pools or reaches, such as those at:
        1. -0.100 km (control)
        2. 0.000 km (control)
        3. 1.800 km (fertilized)
        4. 2.400 km (fertilized)
        5. 2.700 km (fertilized)
        6. 3.500 km (fertilized)
      2. Adults are sampled by angling throughout both reaches; generally, the fertilized zone is sampled between the lowest (furthest downstream) and middle weirs, while the control zone is sampled between the middle and upper (furthest upstream) weirs.
      3. Weirs in Oks Creek are traditionally deployed at -1.000 km, -0.010 km, and 1.650 km.
  3. Sample sites in all other New Reach streams will be determined at the beginning of the year in which they are originally sampled.

Stream discharge

  1. In each stream, discharge will be measured manually at least 6 times (preferably 8 times) per summer at intervals of approximately one week or following significant changes in stream levels. The purpose of these measurements is to create a water level/discharge curve so that the datalogger water level data can be converted to discharge data.
    1. Read and record the river depth on the staff gauge on the side of the stilling well. The units are in hundredths of feet. Record the time of day as well.
    2. At the designated site in each stream (see Section I above), extend a meter tape, perpendicular to the flow, across the river and secure it on both banks. The tape should be relatively taut. This will serve as a reference for doing the transect.
    3. Measure the width of the stream and divide the width into 20 increments. Write these increments in one column of the notebook.
    4. Beginning at one bank, measure and record the depth of the river, then measure and record the current velocity using the Gurley and/or Marsh-McBirney current meter. The meter should be at 60% of stream depth when the current velocity is recorded, i.e., 60% of the way from the surface to the bottom.
    5. Move along the tape to the next increment and repeat the depth and current measurements. Continue until the entire river breadth has been measured. (If the stream bottom is very inconsistent, you may have to take measurements at greater or less than the designated increment along the tape; be sure to record these discrepancies in the notebook). Remove the tape after the transect.
    6. Read the water depth again off of the staff gauge at the stilling well. The average of the depths before and after the discharge measurement is considered the river level for the given discharge.
    7. The discharge is the sum of the products of each individual measurement (an individual measurement is the water velocity at a given increment times the depth at that increment times the distance from the previous increment). This can be computed easily in a spreadsheet.
  2. Water level will be recorded continuously in two ways: 1) with a pressure-sensing probe hooked up to a Campbell Scientific CR-10 datalogger, and 2) with a Stevens float-and-pulley water level chart recorder. The CR-10 datalogger should have 8 fresh D batteries installed at the beginning of every summer; these should last for the entire summer. Stevens recorders are deployed in stilling wells upstream of the pipeline in the Kuparuk and downstream of the Dalton Highway crossing at Oks Creek. A CR-10 will be deployed in Oks Creek just downstream of the stilling well. Doug Kane's group (UA-Fairbanks) will deploy a CR-10 in the stilling well at the Kuparuk. Another CR-10 will be deployed in the New Reach at a designated site. The Stevens Chart recorders are the property of the University of Alaska-Fairbanks (Kuparuk) and the USGS (Oks Creek). New Reach streams will only have CR-10's. A conductivity probe will also be placed in the new stream reach each year.
    1. Program the CR-10 datalogger for each parameter using the owner's manuals. FIRST MAKE SURE THE TIME IS SET PROPERLY ON THE DATALOGGER. The LTER computer has a directory called PC208. Use the program EDLOG for program writing and editing. Transfer the program to the CR10 keyboard and which can then be downloaded to the datalogger. The CR10 Prompt Sheet is vital for using the CR10 keyboard and should be regularly consulted. Decide with Bruce Peterson the number of times per day the datalogger should record river depth. Consult the manual for each probe on how to wire the probes to the datalogger.
    2. Deploy the probes as soon as possible after arriving at Toolik.
      1. Kane's group will deploy the Kuparuk probes.
      2. At Oks Creek, place the datalogger inside the yellow weatherproof metal case (has a hole in the bottom) and their cables should be run through the hole. Place waterproof clay in the hole to surround the cable. Using plenty of cable ties, mount the metal case on the wooden frame supported by rebar (about 5 m downstream from the stilling well). Drive a piece of rebar into the stream bottom in at least waist-high water. Loosely attach a cable tie to the probe cables, loop the tie around the rebar, and slide the probes down so that they rest on the bottom. Place some large rocks on the cables so that the probes stay on the bottom and are somewhat protected.
      3. Locations in new reach streams will be determined after arrival at Toolik.
    3. Data will be downloaded into a storage module for transfer to the LTER computer at the Toolik Camp (both as an ASCII file and on a spreadsheet). You must bring the CR10 keyboard in order to do this Data should be downloaded and backed up weekly, especially in times of potential flooding. The datalogger should be removed if flooding is imminent.
    4. Back at camp, you must use the 9-pin SC532 interface cable attached to the RS232 interface module to communicate between the storage module and computer. Use the SMCOM program (in PC208 directory) from DOS prompt to dump data. Select COM2 as the interface port. Select U for uncollected data and C for comma delineated file. This creates a *.DAT file which can then be imported into a spreadsheet. Keep the *.DAT files as backups. Place the *.DAT files in the directory set up for the specific site. Use the following template for naming downloaded files: YYCJULFL.dat where YY=year, C = site code, JUL is Julian day on which downloaded, and FL = file list e.g. 94N22401.dat is the datalogger file downloaded on Julian Day 224 for the new reach (Blueberry Cr.) in 1994.
  3. Stream height should also be recorded manually each day (if possible) by reading the river depth on the staff gauge and recording it, along with the time and date, in the notebook stored in the stilling well (and on the chart recorder in the stilling well box; write the time, date, and river level on the recorder paper and draw an arrow to the spot at which the chart recorder needle is currently located). These data are used to calibrate the manual discharge measurements with the river height data from the datalogger.
  4. At the end of the season, construct a discharge curve for the stream.
    1. Convert the stage heights from the manual discharge measurements (i.e., the measurements taken in part A of this section) from feet (the measurements taken from the staff gauge) to meters (the depth measurements recorded by the datalogger at the same time and date as the manual discharge measurements).
    2. In Excel (5.0 or higher), plot the 6 (or more) discharge measurements from part A of this section: stage height in meters on the x-axis, discharge in cubic meters per second on the y-axis.
    3. Use the trendline function to draw the line that best fits the discharge data. You should select for a power curve and select for the equation and r2 value to be displayed. (Hint: Be sure to display the numbers in the equation to several decimal places so that your computations in the next steps are precise.)
    4. The equation produced by the relationship will probably be in the form of: y = a(xb), where y is discharge in m3/s, a and b are coefficients, and x is stage height in m.
    5. Now you can calculate the discharge over the course of the season, simply by plugging in the depth readings from the datalogger measurements as the x variable in the equation. Create a discharge column in the spreadsheet containing the datalogger depth measurements, input the formula using the equation (substituting the depth measurements for x) Then, plot the discharge value against the date. This will give you a summer discharge profile.

Temperature

  1. Water temperature will be measured manually each day (if possible) using a 12" blunt stem rheotemp thermometer. The purpose of these measurements is to calibrate the temperatures recorded by the Campbell Scientific CR-10 dataloggers.
    1. In the Kuparuk and Oks Creek, temperature should be measured at the stilling wells.
    2. Hold the thermometer in the water for about 15 s before reading; if possible, keep the bulb in the water when reading the temperature.
    3. Record the day, time, and water temperature in the notebooks that are stored in the stilling wells.
    4. Store the thermometer in the stilling well box.
  2. Water temperature will be continuously recorded by the Campbell Scientific CR-10 datalogger and a Campbell Scientific 107B Temperature Probe. Decide with Bruce Peterson the number of times per day the datalogger should record river temperature.
    1. Program the datalogger using the owner's manual for the CR-10. Use a computer to program the datalogger so that comments and footnotes can be written regarding programming. Instruction 11 will be the important operation for programming the datalogger (see the probe owner's manual).
    2. Deploy the probes simultaneously with the depth sensor probes, and download temperature data along with other datalogger information (see Section II.B).

Fertilization of the stream reaches

  1. Fertilization of the Kuparuk River is done via continuous dripping at a designated drip site. Fertilization of Oks Creek is done via continuous dripping at river km 0.0 (1991-1996). The set up and dripping procedures follow; the fertilization protocol in a given year at a given stream varies and is decided by the Streams PI's on the Arctic LTER project.
  2. Fertilization with ammonium sulfate (NH4SO4) is done to determine the effects of added nitrogen on stream productivity.
    1. Materials:
      1. 55-gallon drums (or 150-gal)
      2. 50-lb bags NH4SO4
      3. Cole-Parmer Masterflex peristaltic pump
      4. Materflex pump drive tubing
      5. 2 1000-ml pipette tips
      6. trolling motor
      7. rebar (4 rods per dripper)
      8. rebar pounder, if rebar is not already in place
      9. rope (with tygon tubing and funnel already attached; the long rope is for the Kuparuk, the short is for Oks Creek)
      10. tygon tubing
      11. short section of tygon tubing threaded through PVC pipe
      12. duct tape
      13. 12-V battery
      14. solar panel
      15. graduated cylinder
      16. stopwatch
    2. Set up: At the designated dripper site, drive two pieces of rebar into the bank on either side of the river. They should be in a straight line perpendicular to stream flow. On each bank, drive one piece straight into the ground (plumb), about a meter from the bank, and another at a 45 angle (away from the river) about three meters from the bank. The angled piece will serve as a deadman.
    3. Secure the funnel to the top of the plumb rebar on the same bank as the dripper setup. Use both the rope (clove hitches or half hitches work well) and the duct tape. Tie the end of the rope securely to the base of the deadman; the rope should be taut.
    4. Extend the rope across the river. Secure it to the top of the plumb rebar on the opposite bank and then tie it off to the base of the deadman. The rope should be a meter or more above the river level, so that rising water does not take the rope out, and the end of the tygon should hang over the center of the channel.
    5. Along the river bank at the designated site, set a 55-gallon plastic drum on solid, level ground. Empty two 50-lb bags of NH4SO4 into the drum.
    6. Pump river water into the drum until the drum is full.
    7. Lower a trolling motor into the drum and run the motor for 10 minutes. This should dissolve the NH4SO4 into solution. (Be careful initially-it is not good for the prop to stick it into big chunks of fertilizer and then run it too hard.) If the trolling motor is not working well, you will have to stir the solution (e.g., with a piece of rebar or an oar) until all of the fertilizer is dissolved.
    8. Hook up a Masterflex peristaltic pump to a 12-V battery and a solar panel in parallel. These should be checked regularly to ensure good connections and a charged battery. The pump should have a short length of drive tubing threaded through the pump head (see Operator's Manual if you need help with this).
    9. Run a length of tygon tubing (without holes) from the funnel to the pump. Connect the tygon to the outlet end of the drive tubing with a pipette tip. Then, connect the length of tygon tubing that is threaded through the PVC pipe to the intake end of the drive tubing with a pipette tip and run the tygon into the drum, all the way to the bottom. When the pump is engaged, the fertilizer should be pumped into the funnel end of the dripper tubing and thus be dripped into the river.
    10. Check the rate of dripping with a graduated cylinder and a stopwatch to be certain that the NH4SO4 fertilizer is being dripped at the proper rate. The target rate for Oks Creek is 148 ml/min.
    11. The volume of fertilizer in the drum should be checked regularly. (The Oks Creek dripper uses 55 gallons of fertilizer each day.)
  3. Fertilization with phosphoric acid (H3PO4) is done to determine the effects of added phosphorus on stream productivity.
    1. Materials:
      1. 180-lb. carboy of H3PO4
      2. Microperpex peristaltic pump
      3. drive tubing
      4. 2 100-ml pipette tips
      5. tygon tubing
      6. short section of tygon tubing threaded through PVC pipe
      7. rebar (4 rods per dripper)
      8. rebar pounder, if rebar is not already in place
      9. rope (with tygon tubing and funnel attached; the long rope is for the Kuparuk, the short rope is for Oks Creek)
      10. 12-V battery
      11. solar panel
      12. graduated cylinder
      13. stopwatch
    2. Refer to the setup steps of Part B of this section for installation of the dripper apparatus.
    3. Along the river bank at the designated site, set a carboy containing 180 lbs. of H3PO4 on solid, flat ground.
    4. Hook up a Microperpex peristaltic pump to a 12-V battery. Connect the posts of the battery to a solar panel (battery and panel in parallel). These should be checked regularly to ensure good connections and a charged battery. The pump should have a short length of drive tubing threaded through the pump head (see pump Operator's Manual if you need help).
    5. Run a length of tygon tubing (without holes) from the funnel to the outlet end of the drive tubing. Connect the tygon to the drive tubing with a pipette tip. Connect the piece of tygon threaded through the PVC to the intake end of the drive tubing with a pipette tip. Run the tygon/PVC into the carboy, all the way to the bottom.
    6. When the pump is engaged, the fertilizer should be pumped into the funnel end of the dripper tubing and thus be dripped into the river (it will take several minutes for the fertilizer to reach the funnel; these pumps are rather slow; turn the pump up to maximum speed (99) until the fertilizer is actually dripping into the river, and then gradually slow it down).
    7. Check the rate of dripping with a graduated cylinder and a stopwatch to be certain that the H3PO4 fertilizer is being dripped at the proper rate. The target rate for the Kuparuk River is ~2.4 ml/min (or 0.32 mol/L of river water, based on a mean summer discharge of 2.0 m3/s). The target rate for Oks Creek is ~1.3 ml/min (or 0.32 mol/L of river water, based on a mean summer discharge of 1.0 m3/s).
    8. The volume of fertilizer in the carboys should be checked regularly to make sure that the carboys have not run dry. They will need to be replaced once in Oks Creek (two total per summer) and twice in the Kuparuk (three total per summer).
  4. 15N Addition
    1. NH4Cl with 10% 15N is added to stream reaches as a label to trace the movement of nitrogen through the food web of the stream.
    2. Materials:
      1. carboy
      2. NH4Cl with 10% as 15N (amount dependent on discharge; see calculations)
      3. Microperpex peristaltic pump
      4. drive tubing
      5. tygon tubing
      6. funnel
      7. 12-V battery
      8. solar panel
      9. graduated cylinder
      10. stopwatch
    3. Refer to the setup steps of Part B of this section for installation of the dripper apparatus.
    4. Along the river bank at the designated site, set a carboy containing 15NH4Cl acidified to pH 3.
    5. Hook up a Microperpex peristaltic pump to a 12-V battery. Connect the posts of the battery to a solar panel (battery and panel in parallel). These should be checked regularly to ensure good connections and a charged battery. The pump should have a short length of drive tubing threaded through the pump head (see pump Operator's Manual if you need help).
    6. Run a length of tygon tubing from the funnel to the outlet end of the drive tubing. Connect the tygon to the drive tubing with a pipette tip. Connect the piece of tygon threaded through the PVC to the intake end of the drive tubing with a pipette tip. Run the tygon/PVC into the carboy, all the way to the bottom. Attach a pipet tip to the end of the tygon tubing by the funnel. Insert tip into the funnel. Funnel should be attached to tygon which is extended to the middle of the stream where the 15N will be dripped in.
    7. When the pump is engaged, the 15N should be pumped into the funnel end of the dripper tubing and thus be dripped into the river (it will take several minutes for the 15N to reach the funnel; the drip rate may be low; so to get it through the tubes faster turn the pump up to maximum speed (99) until the fertilizer is actually dripping into the river, and slow it down just be for).
    8. Check the rate of dripping with a graduated cylinder and a stopwatch to be certain that the 15N label is being dripped at the appropriate rate (ml / min).
    9. The volume of 15NH4Cl in the carboy should be checked regularly; it will not need to be refilled as often as the other two types of fertilizer containers.

Water chemistry and nutrient sampling

  1. Water chemistry and nutrient concentrations in the streams will be measured weekly via transects covering the experimental reaches of each stream. Each sampling station should be clearly marked with stakes at the beginning of the season.
  2. pH and water temperature
    1. Materials:
      1. pH meter
      2. calibration solutions (pH 7.00 and 4.01)
    2. Before going into the field, calibrate the portable pH meter, using the instructions provided with the meter (if you have questions, George Kling is very knowledgeable about the proper calibration and use of these meters).
    3. Using the pH meter, measure water temperature and the pH at each site according to the instructions in the manual. Record the values measured at each site in a field notebook. (Note: it is a good idea to put the pH probe into the water at the beginning of the sampling at each site; the probe takes time to equilibrate, and you can get a lot done while it's equilibrating.)
  3. Conductivity
    1. Materials:
      1. conductivity meter
      2. pH meter (with temperature-measuring capability)
    2. At each station, determine the water temperature with the pH meter (see above). Adjust the manual temperature calibration dial on the face of the conductivity meter to the appropriate temperature.
    3. Switch the conductivity meter to S/cm. Measure the conductivity of the water and record the value.
  4. Sestonic chlorophyll concentration
    1. Materials:
      1. pre-labeled 1-L Nalgene amber bottle
      2. cooler with ice
    2. At each station, take a 1-L Nalgene amber bottle (labeled by river and station) and go to the center of the river. Rinse the bottle 3x by filling the bottle half full with river water, capping the bottle, inverting the bottle several times and shaking vigorously, and then pouring the water out.
    3. After the third rinse, fill the bottle to the top with river water, cap the bottle, and store for return to the Toolik Camp. Repeat this procedure at stations designated for duplicate samples.
    4. When you return to the truck after collecting all of the samples, place the bottles in a cooler with ice to keep them cold until you return to camp.
    5. Upon return to the camp, place the bottles in the refrigerator until ready for analysis. These samples will be used for laboratory measurements of total chlorophyll.
    6. Remember NOT to acid-wash chlorophyll bottles between uses; acid rapidly degrades chlorophyll, so traces of acid from the acid bath could affect the chlorophyll in the samples.
  5. Nutrients, sestonic particulates, and ions
    1. Dissolved nutrients and sestonic particulates will be collected at every station. Ions (cations, anions, and alkalinity) will only be collected from a single control station and a single fertilized station in each stream (Kuparuk, .564 km and 2.065 km; Oks Creek, -.136 km and 1.054 km; New Reach streams to be determined).
    2. All water samples will be collected and filtered in the field according to the following protocol.
    3. Materials:
      1. 60cc sterile syringes
      2. pre-combusted 25-mm GF/F filters
      3. 25-mm Gelman filter pre-labeled Gelman petri dishes-L plastic filter flask
      4. tweezers
    4. Place on 25-mm GF/F filter into each acid washed filter cassette. This can be done in the lab to save time and avoid contamination in the field. Handle filters with the tweezers only.
    5. At each station, go to the center of the stream and rinse a 60cc syringe 3 times with stream water.
    6. Fill the syringe with stream water, mount the filter cassette (with filter) onto the syringe, and fiilter water into the other sample bottles as described in the steps to follow. When you run out of water, put the second filter cassette (with filter) onto the syringe and filter a second 500-ml sample, again from the center of the river. There is no need to rinse the filter cassette or syringe a second time through. A second filter cassette may already be set up to save time (2 per station). This water can also be used to fill the bottles in the steps below.
    7. After the last water has gone through the filter, remove it with the tweezers and place it in a petri dish labeled for particulates (see below, Part H). For each transect place one filter in a petri dish that has not been used for filtration. This will be used as the blank. Do this for both for PN/PC and PP.
    8. Avoid making any physical contact with the filtered water! Your skin can contaminate the water for certain analyses, especially ammonium.
  6. Nutrients (NH4+, PO4-3, and NO3-)
    1. Materials:
      1. 50-ml aliquot of water from the filter flask (see Part E above), plus extra water to rinse the bottle
      2. 60-ml HDPE bottle, pre-labeled, acid washed and rinsed with DI (may be designated to a station and reused on every transect).
    2. Filter a small amount of water into the bottle, cap it, and shake it vigorously to rinse it. Pour the water out. Repeat twice.
    3. Fill bottles with 50-60 ml of filtered stream water. Cap securely and store it for return to Toolik.
    4. Upon return to camp, if the samples are not to be run immediately, store the bottles in the refrigerator.
    5. Ammonium will always be analyzed manually in New Reach streams where the 15N ammonium is being added. A water sample for ammonium hand chemistry should be collected in the manner described below.
      1. Materials:
        1. 20-ml aliquot of water from the filter flask (see Part E above), plus extra water to rinse the centrifuge tube
        2. clean (prereacted - see manual chemistry protocol), dry pre-labeled 56-ml centrifuge tube
        3. 250-ml HDPE bottle (optional; see below)
      2. Pour a small amount of filtered water from the filter flask into the centrifuge tube, cap it, and shake it vigorously to rinse it. Pour the water out. Repeat twice.
      3. Pour 20 ml of filtered water into the centrifuge tube. Cap it and store it for return to Toolik.
      4. Note: if the autoanalyzer is not operating reliably, then pour at least 200 ml of filtered water into a 250-ml HDPE bottle. Return to camp and distribute 20 ml of the filtrate water into three 50-ml VWR tubes and refrigerate. Consult the autoanalyzer technician for manual analytical methods for nitrate and phosphate.
      5. Upon return to camp, store the bottles in the refrigerator until they are ready to be analyzed.
  7. Total dissolved nitrogen and phosphorous (TDN and TDP)
    1. Materials:
      1. 50-ml aliquot of filtered stream water from Part E (above)
      2. 60-ml HDPE bottle, pre-labeled for TDN and TDP samples
    2. Pour about 50 ml of filtered water into the each bottle. Cap securely and store for return to camp.
    3. Upon return to camp, preserve w/ acid (use HCl). at 1ul of 6N acid per ml of sample.
    4. Place sample in thick freezer ziploc plastic bags and seal. The bags will reduce NH3 adsorption by the sample. Store in the refrigerator.
    5. Ship samples in coolers to MBL at the end of the field season.
  8. Sestonic particulates (PN/PC and PP)
    1. Materials:
      1. two pre-ashed GF/F filters that have each had 500 ml of stream water filtered through them (see Part E above)
      2. two 25-mm Gelman filter cassettes
      3. two clean, dry, pre-labeled petri dishes
      4. tweezers
    2. After each filter has had 500 ml of water run through it, remove the filter with the tweezers and place it into one of the petri dishes. The dishes should be pre-labeled (stream, station, date, and either PNPC, particulate carbon and particulate nitrogen, or PP, particulate phosphorus; since the two filters at a given station will be identical, they can go into either the PNPC dish or the PP dish).
    3. After returning to the lab, immediately place these filters into the drying oven at 50 degrees C for 24 - 48 hours.
    4. Remember to dry field filter blanks for each type of particulate sample (1/transect).
    5. Once the filters are dry, store them in sealed plastic bags and ship to MBL.
  9. Cations
    1. Materials
      1. 50-ml aliquot of water from the filter flask (see Part E above)
      2. clean, dry pre-labeled 60-ml HDPE bottle
    2. Cations will be collected at only two stations per stream: one control and one enriched. In the Kuparuk, stations 0.564 km(control) and 2.065 km(fertilized). In Oks Creek, the stations are -0.136 km(control) and 1.056 km(fertilized). In New Reach streams, the stations will be determined at the beginning of the field season.
    3. Pour about 50 ml of filtered water into the cation bottle. Cap securely and store for return to camp.
  10. Anions
    1. Materials
      1. 30-ml aliquot of water from the filter flask (see Part E above)
      2. clean, dry pre-labeled 30-ml HDPE bottle
    2. Anions will be collected at only two stations per stream: one control and one enriched. In the Kuparuk, stations 0.564 km(control) and 2.065 km(fertilized). In Oks Creek, the stations are -0.136 km(control) and 1.056 km(fertilized). In New Reach streams, the stations will be determined at the beginning of the field season.
    3. Pour about 30 ml of filtered water into the anion bottle. Cap securely and store for return to camp.
  11. Alkalinity
    1. Materials
      1. about 150 ml of water from the filter flask (see Part E above)
      2. pre-labeled 125-ml HDPE bottle (may be designated and reused during each transect)
    2. Alkalinity will be collected at only two stations per stream: one control and one enriched. In the Kuparuk, stations 0.564 km(control) and 2.065 km(fertilized). In Oks Creek, the stations are -0.136 km(control) and 1.056 km(fertilized). In New Reach streams, the stations will be determined at the beginning of the field season.
    3. Use about 20-30 ml of filtered stream water to rinse out the urine cup; pour the water into the cup, cap it, shake vigorously, and pour out.
    4. Pour about 120 ml of filtered water into the urine cup, cap it securely, and store for return to camp.

Dissolved inorganic carbon (DIC)- in progress

Dissolved organic carbon (DOC)

  1. Materials
    1. 20-ml aliquot of water from the filter flask (see Part E above)
    2. clean, dry pre-labeled 20-ml glass scint. vial
  2. DOC will be collected at only two stations per stream: one control and one enriched. In the Kuparuk, stations 0.564 km(control) and 2.065 km(fertilized). In Oks Creek, the stations are -0.136 km(control) and 1.056 km(fertilized). In New Reach streams, the stations will be determined at the beginning of the field season.
  3. Fill vial with 20-ml of filtered water. Cap securely and store for return to camp.

2 x 2 Epilithic algal scrubs

  1. Algae growing on the stream bottom rocks are an important component of primary productivity. Therefore, samples of epilithic algae are scrubbed off of the rocks and their chlorophyll content is measured. Particulate nutrients are measured as well.
  2. The following methods are based on methods from the LTER database, previous protocol sheets, and Peterson et al. (1993), and have been modified into outline form.
    1. Materials
      1. wash basin
      2. steel bristle scrub brush
      3. 56-ml centrifuge tubes (5 per station)
      4. 250-ml squirt bottle
      5. 2x2 slide holders
      6. funnel
    2. At each station, rinse the wash basin, scrub brush, slide holders, and squirt bottle 3X with river water. Fill the squirt bottle.
    3. From a riffle, select 5 rocks that fit the following criteria:
      1. rocks with no filamentous algae or moss (to eliminate overestimates of chl due to filamentous algae or moss)
      2. rocks with fairly smooth upper surface (uneven surfaces prevent efficient removal of epilithon)
      3. rocks that have been submerged for a long period of time.
    4. It is possible that very few rocks at some sites will meet all of the above criteria. If you must select rocks that do not fit any or all of the criteria, make careful and thorough notes describing the deviations.
    5. Place the slide holder over a smooth portion of the upper surface of the rock. With the brush, scrub the area within slide holder. Hold the rock over the basin so that all scrubbate falls into the basin.
    6. With the squirt bottle, rinse the scrubbed area, the holder, and the brush into the basin. DO NOT SQUIRT MORE THAN 56 ml INTO THE BASIN.
    7. Pour the contents of the basin into a labeled (by river and station) centrifuge tube. Use the funnel to facilitate pouring.
    8. Bring volume in the centrifuge tube up to the rim (i.e., to 56 ml) with river water (from squirt bottle)
    9. Repeat steps 2-9 for each rock at each station.

Whole rock scrubs

  1. The 2 x 2 rock scrubs outlined above are a fast method used to determine the chlorophyll and particulate nutrients in the epilithic algae. Whole rock scrubs are a more thorough method for the same purpose. Once a summer, whole rock scrubs will be conducted at the same time as 2 x 2 scrubs to compare the results and efficiencies of the two methods.
  2. The methods below are those of Bruce Peterson (personal communication) and have been modified into outline form.
    1. Materials
      1. wash basin
      2. steel bristle scrub brush
      3. 1-L plastic graduated cylinder
      4. 250-ml squirt bottle
      5. 100-ml plastic bottle, pre-labeled
    2. One time late in the summer, after the chlorophyll has had a chance to build up on the stream bottom, on the same day as a 2 x 2 scrub transect, conduct a whole-rock scrub transect as well. Whole rock scrubs will be done at three of the same sites as 2 x 2's: one in the control reach, one in the fert zone close to the dripper, and one elsewhere in the fert zone that typically has high chlorophyll. (Note: this does not mean that 2 x 2's will not be done at these three stations!)
    3. At each site where whole rock scrubs are to be done, rinse the wash basin, graduated cylinder, brush, and squirt bottle 3x with river water.
    4. Select several rocks from each station that fit the following criteria (make careful notes of any rocks that do not fit the criteria):
      1. rocks with no obvious filamentous algae or moss (to avoid overestimates of chl due to filamentous algae or moss)
      2. rocks with fairly smooth surface (uneven surfaces prevent efficient removal of epilithon)
    5. The rocks should be of appropriate size to form a single layer of rocks that fills the bottom of the wash basin. Remove the rocks from the basin and gently set them aside, right-side up.
    6. Fill the graduated cylinder with 1 liter of stream water. Pour ~250 ml into the squirt bottle and pour the rest into the basin. All of this water will be needed, so do not spill or waste any.
    7. One by one, scrub each rock vigorously and thoroughly with the wire scrub brush. All scrubbate should fall into the basin.
    8. Rinse each rock with the water from the squirt bottle, making sure that all rinse lands in the basin (be sure to save enough water in the squirt bottle to rinse all of the rocks and the scrub brush).
    9. After all scrubbing is completed, the basin will contain a slurry of 1 liter of water plus all of the scrubbate.
    10. Stir up the slurry in the basin so that it is homogenous; then fill the pre-labeled 100-ml bottle with slurry. This is a subsample for laboratory analysis.
    11. Pour out the remaining slurry. Store the bottle for return to camp.
    12. Repeat this procedure twice, so that at each of the three stations, you scrub a total of three basins of rocks.

Epilithic primary productivity

  1. In addition to the studies involving the rock scrubs (see above, Sections VI. and VII.), rocks will be incubated in special laboratory chambers to determine primary productivity of the epilithic algal layer. The amount of photosynthesis and respiration occurring on individual rocks will be determined during the incubation period. These rocks may have filamentous algae; the rock scrubs excluded such rocks when possible.
  2. The methods below are based on methods written by Breck Bowden, the PI for this project, in the document files of the Arctic LTER database in 1989 and 1990 (filenames 89BOMETA.DOC and 90BOMETA.DOC). They have been modified into outline form.
    1. Materials:
      1. carboy (optional)
      2. coolers
    2. At the scheduled time (determined at the beginning of the field season), collect rocks at random locations within pools or riffles at each sample site (locations vary from year to year). Select surface rocks with a "typical" development of epilithon, based on visual inspection of the site, for study. Reject rocks with heavy moss growth and instead choose rocks of a uniform size and shape, that would fit neatly in the chamber bottom.
    3. In the Kuparuk River, 3 to 4 rocks of a modal shape and size essentially covers the chamber bottom in a single layer, at a surface density similar to that found in the river. Substrate size in Oksrukuyik Creek is substantially smaller than in the Kuparuk River; thus, use 4 to 7 rocks from Oksrukuyik Creek to cover each chamber bottom. (Substrate sizes for New Reach streams will be determined at the beginning of the summer.)
    4. Collect water for the incubations in carboys (optional; Toolik Lake water may be used instead).
    5. Place rocks collected from the river in coolers, without water, to keep them cool and moist. (If the rocks are kept submerged in water in the coolers, delicate epilithic material will be dislodged during transport from the field to the lab. Without water, the rocks and epilithon can be transported with the epilithon essentially intact, even with flocculent pool epilithon.) The time from collection and transport to installation in a chamber is generally 1-2 h, and should be minimized.
    6. Upon return to Toolik, immediately place rocks and water into the experimental chambers in Bowden's polar tent.

Bryophytes

  1. When the fertilization of the stream reaches began, the response of bryophytes (mosses and liverworts) was not anticipated or examined. However, bryophytes have increased dramatically in the fertilized reaches. Thus, the amount of stream bottom covered by bryophytes is now studied.
  2. The methods below are based on methods written by Breck Bowden in the document files of the Arctic LTER database (files titled BOMOSS.DOC) and have been modified into outline form.
    1. Materials
      1. meter tape
      2. plexiglas viewscope
    2. Five transects will be conducted at each designated riffle site; sites will be selected early in the field season.
    3. Extend the meter tape across the width of the stream.
    4. For each transect, note cover along entire transect by proceeding at 5-, 10-, or 20-cm intervals across width of stream. Use the plexiglas viewscope to maximize visibility; hold the scope concave side up with the "lens" just below the surface of the water.
    5. At each interval, note the presence of epiplithic bryophytes or macroalgae beneath the target point on the viewscope.
    6. If the point appears bare, note "bare" (bare rocks have epilithic diatom communities too small to be seen by the unaided eye). If bryophytes or macroalgae are present, note the predominating species present (you will only write one species for any particular point; this is a problem when there is algae growing on a bryophyte).
    7. The percent cover of each species is "% of total observations and estimating area." Percent cover estimates are expressed as averages of all transects (n=5) at each riffle site

Bioassays of epilithic algae

  1. Bioassays of epilithic algae are done using small-scale artificial nutrient-diffusible substrata. These bioassays produce many replicates of numerous treatments at minimal cost and time expenditure.
  2. The methods below are based on Gibeau and Miller (1989) and have been modified into outline form. Mike Miller is the PI for this project.
    1. Materials:
      1. agar vials (see below)
      2. porous porcelain discs, soaked in HCl (see below)
      3. wooden vial holders (see below)
      4. silicon sealant
      5. rope to secure vial holders (should be several meters longer than width of stream)
      6. 2 metal spikes per wooden vial holder
      7. pliers (bring when time to remove vials from river)
      8. 125-ml urine cups (one per vial) (bring when time to remove vials from river)
    2. Three assay experiments will be conducted in each stream during each summer. The dates and sites will be determined at the beginning of the field season. Each experiment lasts three weeks.
    3. Each experimental chamber is composed of a 10-dram plastic vial (Dynalab Corp. #2636-0010) used as a reservoir, filled with various nutrient-supplemented agar treatments.
    4. The agar treatments are 37-ml of a 2% (w/v) Difco Ultrapure Agar solution augmented with one each of the following treatments:
      1. control (plain agar)
      2. humic acid extract plus phosphorus (2 g humics/L and 0.5 ml conc. HCl plus 0.005 moles K2PO4/L)
      3. phosphorus (0.005 moles K2PO4/L)
      4. ammonia (0.05 moles NH4Cl/L)
      5. phosphorus plus ammonia (0.005 moles K2PO4/L + 0.05 moles NH4Cl/L)
      6. vitamins (B1 0.1 mg/L, plus Biotin 5 mg/L)
      7. a trace metal mixture (Woods Hole formula plus 0.0999g NTA/500 ml as a chelator; Stein 1973).
    5. All agar treatments should be autoclaved to ensure sterility. (Does this affect the vitamin treatment?)
    6. The chamber is sealed with a coarse, porous porcelain or fused silica (2.6-cm diameter) disk (crucible cover) (Leco Corp. #528-041) that has been cleaned by soaking in a 10% HCl for 48 hr and rinsing copiously with distilled water.
    7. Heat each disc on a hot plate until hot enough to melt the plastic (?). Seal the agar-filled vial by placing the hot disc on top of the vial, melting the plastic at the mouth of the vial, and molding it around the disc.
    8. Turn the vial upside-down, allowing the agar mixture to solidify in contact with the porous disc.
    9. Cap and color-code finished vials according to the treatment they contain.
    10. Arrange the vials in batches of 42; each batch will contain 6 replicates of each of the seven treatments.
    11. Place each batch in a wooden holder(s), which are strips of lumber with pre-drilled holes (3-cm diameter) and mounted on 1.2 m x 0.31 m plywood. Secure the vials into the holes with a small spot of silicon sealant on the bottom of each vial (use as little as possible to avoid the effects of acetic acid leaching from the sealant).
    12. At each site in the stream, secure the wooden holder to the stream bottom. Use two restraints: a rope running between opposite shores and looped through a hole on the upstream side of the wooden base; and two metal spikes at each end of the board, driven through the base and into the rocky bottom, with flat rocks placed over the stakes at each end. This should ensure that the boards will remain stationary on the river bottom even in high flow periods.
    13. Leave the boards and the agar vials undisturbed for three weeks. This procedure will be done three times, with one week of overlap between experiments.
    14. When an incubation period is over, unfasten the board from the river bottom but keep it submerged. Carefully maneuver it to the shore.
    15. Remove the vials one at a time by color code and place the discs into pre-labeled plastic 125-ml urine cups. The discs can be removed by gently squeezing the mouth of the vial with pliers.
    16. Three discs from each treatment can be placed in the cups dry; they will be assayed for chlorophyll biomass. The other three should be placed in cups with about 50 ml of water and remain completely submerged; they will be assayed for primary productivity.
    17. Return the samples to the lab and give them to Miller's group.

Insects

  1. To estimate density, growth, and production of stream insects during the summer, surveys of stream insects are done using either a drift sampling technique or a rock-scrubbing technique.
  2. The methods below are those of Anne Hershey, the PI for this project, and have been modified from LTER database document files, Peterson et al. (1993), and Hershey and Hiltner (1988) into outline form.
  3. Drift sampling technique
    1. Materials:
      1. drift net (of known area)
      2. two long rebar stakes, sledgehammer
      3. current meter
      4. meter stick
      5. stopwatch
      6. 100-m sample net
      7. wash basin
      8. 6" funnel
      9. 250-ml widemouth sample jars
      10. 95% ethanol
      11. labels
    2. At a riffle at each sampling station, pound the rebar stakes into the substrate and use them to anchor the drift net.
    3. Deploy the drift net and record the exact time that the net begins sampling. Begin timing with the stopwatch. Check the net to make sure it is untangled.
    4. After the net is deployed, immediately measure the current at the mouth of the drift net.
    5. With the meter stick, measure the portion of the net's mouth that is above the waterline. The current data, the area of the net's mouth below the waterline, and the total sampling time (in seconds) will be multiplied together to give the total water volume sampled.
    6. Watch the net to make sure that it does not become clogged. At high flows, clogging can occur within 1-2 minutes. At low flows, clogging may not occur for 30 minutes. A 15-minute sample has been used in previous samples. If clogging occurs, then pull the net and record the exact time it was in the river.
    7. When the sampling time is nearly complete, record the current at the mouth of the net again. The current should be approximately equal to the current from the beginning of the sample.
    8. Pull the net and stop the stopwatch. Record the exact time that the sampling was concluded.
    9. Hold the net vertically (with the jar end down) and rinse down the contents of the net into the jar.
    10. Loosen the hose clamp that holds the jar to the drift net. Rinse the drift net jar in the basin with some river water. Pour the contents of the basin through the 100-m sample net.
    11. Transfer the contents of the sample net to the 250-ml widemouth jar. Use the funnel to facilitate the transfer.
    12. Label and preserve the samples in 95% ethanol.
    13. Repeat this procedure, so that there are two samples from each station.
    14. Return the samples to camp for shipment to the University of Minnesota-Duluth, where they will be picked, sorted, counted, and measured using a digitizing pad.
  4. Rock-scrubbing technique
    1. Materials:
      1. plastic basin
      2. soft-bristle scrub brush
      3. 100-m net
      4. 250-ml widemouth jars
      5. 6" funnel
      6. 95% ethanol
      7. labels
    2. At each station, select a riffle habitat, similar in depth and flow to riffles sampled at other stations.
    3. A rock-scrub sample consists of four rocks collected haphazardly from each riffle. Place the rocks in a plastic basin. The rocks should cover ½ to ¾ of the bottom of the basin. The rocks will have an estimated average upper rock surface area of 363 18 cm2. Be certain to transfer the rocks from stream to basin as quickly and carefully as possible to minimize the loss of insects.
    4. Add about 1 liter of water to the basin.
    5. Scrub the rocks with soft nylon-bristle brushes to remove the insects. The insects should fall into the basin.
    6. Pour the insects from the basin through the 100-m mesh net, which will seive out and concentrate the insects.
    7. Transfer the insects into a widemouth jar. Use the funnel to facilitate the transfer. Rinse the net into the jar with 95% ethanol.
    8. Label and preserve the samples in 95% ethanol.
    9. Repeat this procedure, so that there are two samples from each station.
    10. Return the samples to camp for shipment to the University of Minnesota-Duluth, where they will be picked, sorted, counted, and measured using a digitizing pad.

YOY Arctic grayling

  1. Length and weight analysis
    1. Materials:
      1. dipnets (large and small)
      2. backpack electrofisher
      3. orange rubber gloves
      4. labeled 1-L plastic bottles
    2. Before operating the electroshocker, make sure that everyone present is wearing orange rubber gloves for insulation!
    3. Each week, at three sites in the control reaches and three in the fertilized reaches of each stream, (each site should be several hundred meters apart), catch YOY using dip nets or backpack electrofisher. Place YOY in pre-labeled 1-L plastic bottles ¾ full of water (no more than 10 YOY per bottle). Replace water hourly while in the field. Try to get at least 10 YOY from the control zones and 10 from the fertilized zones of each stream.
    4. Return live fish to lab.
    5. See measuring and weighing protocol in lab section.
    6. Hint: early in the season, YOY will be found primarily in slower waters (like pools and backwater channels) in sheltered areas; by August, they move out into faster flows.
  2. Gut content analysis
    1. Materials:
      1. dipnets
      2. backpack electrofisher
      3. orange rubber gloves
      4. labeled 250-ml plastic bottles
      5. 95% ethanol
    2. Before operating the electroshocker, make sure that everyone present is wearing orange rubber gloves for insulation!
    3. Each week, catch YOY using dip nets or backpack electrofisher (described above) and reserve 5 YOY from each site. Transfer these fish to a labeled 250-ml bottle.
    4. Preserve the 5 reserved fish in the field with 95% ethanol.
    5. Return fish samples to lab.
    6. See gut content analysis protocol in lab section.
  3. Otolith marking
    1. Materials:
      1. dipnets
      2. backpack electrofisher
      3. orange rubber gloves
      4. 1-liter plastic bottles
    2. Before operating the electroshocker, make sure that everyone present is wearing orange rubber gloves for insulation!
    3. Early in the season, in each stream, catch at least 10 YOY (5 from control, five from fertilized reach) using dip nets or backpack electrofisher. Place YOY in pre-labeled 1-L plastic bottles ¾ full of water (no more than 10 YOY/bottle). Replace water hourly while in the field.
    4. Return live fish to the lab, where they will be marked and tagged.
    5. See otolith marking protocol in lab section.
    6. Release fish the following day at same site where they were caught.
    7. Electroshock the same location 3-5 days following release. Return any recaptured YOY live to the lab; you will know that they are recaptures by the subcutaneous acrylic paint tags.
    8. Repeat mark recapture (steps 1-7) at a later time (i.e. perform this procedure twice per summer).
  4. Isotopes
    1. Materials:
      1. dipnets
      2. backpack electrofisher
      3. orange rubber gloves
      4. 1-liter plastic bottles
      5. labels
      6. aerators
      7. buckets (or 1-liter bottles)
    2. Before operating the electroshocker, make sure that everyone present is wearing orange rubber gloves for insulation!
    3. Toward the end of the season, in each stream, catch at least 10 YOY (5 from control, five from fertilized reach) using dip nets or backpack electrofisher. Place YOY in pre-labeled 1-L plastic bottles ¾ full of water (no more than 10 YOY/bottle). Replace water hourly while in the field.
    4. Return live fish to camp and place in aerated 1-L bottles or buckets of water.

Adult Arctic grayling

  1. Tagging, length and weight analysis
    1. Materials
      1. fly rods
      2. flies (mainly elk-hair caddis, blue-winged olive, nymphs)
      3. spinning rods
      4. mepps spinner (#0 or #1; larger sizes can injure the eyes of the fish!) with barbless double-hook
      5. small nylon-mesh holding bags
      6. large dip nets
      7. fish holding pens: 4x4x4 nylon-mesh, with 4 rods of rebar per pen to anchor pen in the stream
      8. anesthesia: 100 mg MS-222 (Finquel) per liter of water buffered with sodium bicarbonate (NaHCO3) until pH paper is around pH 7
      9. pH paper
      10. Floy t-bar tags and tag injector
      11. PIT tags, PIT tag reader, PIT tag injector
      12. ethanol
      13. fish measuring board (50-cm)
      14. battery-operated portable field scale
      15. buckets
    2. Early in the season, after the weirs are in place, collect as many fish as possible by angling (occasionally by electrofishing or seining)
    3. Hold fish in pens overnight (to allow guts to clear)
    4. Anesthetize fish using 100 mg MS-222 per liter of water buffered with sodium bicarbonate (NaHCO3).
    5. Sterilize tag gun needles with ethanol.
    6. Tag each fish with both:
      1. individually numbered, color-coded Floy t-bar tags (through the flesh on the left side, just below the dorsal fin)
      2. PIT tags (subcutaneous, anterior of the pelvic fins on the ventral surface)
    7. Measure total length of fish to nearest 0.1 cm (from tip of nose to bottom lobe of caudal fin; if caudal fin is missing or damaged, mention in notebook)
    8. Measure wet weight to nearest g using the portable field scale.
    9. Record the tag number, color, weight, length, and release site (i.e. control or fertilized zone) of each fish in the grayling notebook.
    10. Place the fish in the stream. Hold the fish by the caudal-peduncle, head into the current, until the fish has had an opportunity to recover from the anesthesia. Release the fish. Check with Linda Deegan to see how many fish should be released in the control section and how many should be released in the fertilized section.
    11. At the end of the season, the control and fertilized reaches will be fished again. The tagged fish will be recaptured, remeasured and reweighed (see previous section) to determine the amount of growth that occurred during the season, and then released.
  2. Gut content analysis
    1. Materials for gut content analysis
      1. fly rods
      2. flies (mainly elk-hair caddis, blue-winged olive, nymphs)
      3. spinning rods
      4. mepps spinner (#0 or #1) with barbless double hook
      5. small nylon-mesh holding bags
      6. large dip nets
      7. fish holding pens (see above)
      8. 60-cc syringe
      9. 13-gauge needle tipped with rubber tubing tip
      10. 100-m nylon mesh
      11. 95% ethanol, squirt bottle
      12. catchment pan
      13. funnel
      14. 250-ml plastic bottles
    2. Twice per summer, catch 10 fish in each stream (5 control, 5 fertilized reach) by angling. Weigh and measure the fish (see previous section).
    3. While it is still anesthetized, hold the fish with its mouth over the catchment pan.
    4. Fill a 60-cc syringe (loaded with a 13-gauge needle, tipped with rubber tubing to prevent scratching) with water and carefully insert tubing down the fish's esophagus.
    5. Inject the water, forcing the fish to egest its stomach contents through the mouth and into the pan. Gentle pressure on the fish's stomach helps to induce egestion.
    6. Filter stomach contents through the 100-m nylon mesh.
    7. Transfer the stomach contents from the mesh into a 250-ml plastic bottle (use the funnel to make this easier). Label and preserve stomach contents in 95% ethanol.
    8. Place the fish in the stream. Hold the fish by the caudal-peduncle, head into the current, until the fish has had an opportunity to recover from the anesthesia. Release the fish.
    9. Return stomach contents to camp. Store in a sturdy box and ship or hand-carry to MBL for analysis.
  3. Isotopes and otoliths
    1. Materials
      1. fly rods
      2. flies (mainly elk-hair caddis, blue-winged olive, nymphs)
      3. spinning rods
      4. mepps spinner (#0 or #1) with barbless double-hook
      5. small nylon-mesh holding bags
      6. large dip nets
      7. fish holding pens: 4x4x4 nylon-mesh, with 4 rods of rebar per pen to anchor pen in the stream
      8. anesthesia: 100 mg MS-222 (Finquel) per liter of water buffered with sodium bicarbonate (NaHCO3) until pH paper is around pH 7
      9. pH paper
      10. fish measuring board (50-cm)
      11. battery-operated portable field scale
      12. buckets, labels
    2. Late in the season, collect 6 fish from each stream (3 control, 3 fertilized) by angling (occasionally by electrofishing or seining)
    3. Hold fish in pens overnight (to allow guts to clear)
    4. Anesthetize fish using 100 mg MS-222 per liter of water buffered with sodium bicarbonate (NaHCO3). Weigh and measure the fish (see above). Return the fish to the anesthesia until they are dead. Place the fish in buckets and label (date, stream, section).
    5. Return the fish to camp, and immediately freeze them. Ship frozen to MBL for stable isotope analysis and otolith removal.

Conservative tracer studies

Rationale: The general design of this task is that a known concentration of solute is released at a constant rate into the stream for one to several hours and measurements are made downstream to determine the concentration and timing of the passage of the solute pulse. There are two components of this task. The first involves injection of a conservative solute tracer (Cl or Br) to determine the hydraulic properties of the stream. Chloride has been widely used a conservative tracer in many studies because it is cheap and easily measured. Plan to use Cl unless your background Cl concentrations are high (> 5 mg/L) or unless discharge is so high that you cannot release Cl fast enough to achieve measurable concentrations (approx. 10 mg/L above background concentration). In these cases, plan to use bromide as an alternative. Bromide can be measured at levels about 0.1 below the detection limit of Cl. Both Cl and Br can be measured on site with ion specific electrodes. Chloride can also be measured with a high quality conductivity meter. The injection should be performed using a peristaltic pump. The length of the injection time will vary depending on size and discharge of the stream. In general, the injection should continue until the concentration at the downstream station has reached a constant level (plateau) for at least 0.5 hour. In most streams this will require 1-3 hours. The downstream station should be located 50-300 m downstream from the injection site, again depending on the size and discharge of the stream (this should be the same reach used for the 15N addition, or at least the upper part of the 15N addition reach). Data from the conservative tracer injection will be analyzed using an advection-dispersion model with transient storage.

The second part of this task is conducted along with the conservative tracer injection. The stream is slightly enriched with an inorganic nutrient and uptake of the nutrient is measured assuming 1st-order uptake kinetics, and the nutrient uptake length is calculated. Because nutrients are added at levels above background, this method may overestimate uptake length; however, it will give us relative numbers for comparisons. Injections of NO3, NH4, and PO4 should be done on consecutive days (a conservative tracer must be added with each nutrient injection to correct for dilution, but intensive sampling of the conservative tracer for determination of hydraulic properties need only be done with one of the nutrient injections). Rather than specify one standard addition at all sites, each site should aim for additions that are about 2x to 5x the background levels of the nutrient being added. In choosing the concentration addition level, use the following criteria:

  1. the addition should be a small as possible so as to remain below the saturation level (if at all possible) for uptake if it is the limiting nutrient;
  2. the concentration increase must be able to be accurately measured even after about 2/3 of it has been removed from solution.

Set up a lot of stations within the expected uptake length distance below the injection site at which to collect samples (6-8 stations at a minimum - the more measurements of the nutrient increase relative to conservative tracer over distance, the better the estimate of uptake length given the limits on analytical precision), and collect numerous background samples (at least 4-5) and steady state samples (5-10) at each station. Because there is less carryover effect of NO3, its injection should be done on the first day, then NH4, then PO4.

It would be best to complete the conservative tracer and nutrient injections before the 15N addition begins, although they could be done during the 15N addition (short-term addition of nutrients - even NH4 - should not affect the longer-term 15N dynamics). If conditions change substantially during the 6-week 15N addition, it would be best to repeat the short-term conservative tracer and nutrient injections during the latter part of the 15N addition.

  1. Conservative Tracer Injection Methods: The methods below have been modified from: J. R. Webster and T. P. Ehrman, 1996, Solute dynamics, pages 145-160 in F. R. Hauer and G. A. Lamberti (eds.), Methods in Stream Ecology, Academic Press, San Diego. They are for a 100-m reach and using a Mariotte bottle and Cl specific probe. However, we should have a peristaltic pump at all sites to do the injections, so you can replace any Mariotte bottle instructions with those for a peristaltic pump.
  1. Materials and supplies:
    1. Lab -conservative solute (we use non-iodized table salt)
    2. distilled water
    3. containers for standards -- 8
    4. carboys for stock solution of solutes
    5. graduated cylinders (100 mL and 1000 mL)
    6. Field - Peristaltic pump
    7. velocity meter (optional)
    8. meter stick
    9. stop watches
    10. flagging tape
    11. permanent marking pen
    12. tape measure (50-100 m)
    13. squirt bottles with distilled water -- 2 or 3
    14. thermometer
    15. water resistant paper or notebooks, pencils -- for each Cl probe site and release site
    16. bucket
    17. graduated cylinder (100 mL)
    18. sample bottles -- 20 clean
    19. Cl probes -- 4
  2. Mix stock solution of sodium chloride by dissolving 238 g salt per 1 L distilled water. Total volume needed will depend the duration of releases and release rate. Heating the mixture in a water bath aides in dissolution. Mix vigorously and repeatedly for the solution is close to saturation. Make certain the salt is completely dissolved.
  3. Prepare a series of chloride standards (1-20 mg/L) for calibrating the probes. We use 0.5, 1, 2, 3, 4, 5, 10, and 20 mg/L.
  4. Calculate stream flow and necessary release rate to raise stream concentration appropriately (by about 10-15 mg/L at the upstream station).
  5. Use a tape measure to delimit the extent of the experimental reach. Mark every 5 m within the reach with labeled flagging tape.
  6. At each 5-m cross-section, measure wetted channel width, depth across the stream (every 10-20 cm depending on width, minimum 10 depth measurements per cross-section), and thalweg velocity (optional). It may be better to do these measurements after the releases to avoid unnecessary stream stomping. Stream temperature and gradient should also be measured.
  7. Calibrate the Cl probes with the standards. The standards should be placed in the stream until they equilibrate with ambient stream temperature.
  8. FIELD RELEASE: These releases can be done with 2 experienced people, but with more people, the chances of getting good data are much better. It works well with 5 people: one to do the release, 3 reading Cl probes (downstream and 2 intermediate sites), and one person to "coordinate", i.e., make sure everything is done correctly, cover while someone else takes a break, pass out coffee and donuts, and generally keep everyone else happy. No experience necessary.
  9. Collect a series of background (immediately pre-injection) water samples in mid-stream at 10-m intervals over the reach. Work from downstream up and avoid unnecessary stomping in the stream.
  10. Position chloride probes at 20, 50, and 100 m sites. Place probes securely in a well-mixed areas.
  11. Add solute solution to the container. Refer to section III, part D for pump installation and operation.
  12. Synchronize stop watches and open spigot to commence release.
  13. Frequency of chloride readings at downstream site depends upon rate at which the concentration changes in the stream. Record probe readings every 1-5 min (flow dependent) until pulse arrives and then measure every 15-30 seconds as chloride concentration increases rapidly.
  14. At plateau (10 min to several h after commencing release), working from downstream to upstream, take 1 sample in a clean bottle from mid-stream at 10-m intervals for Cl analysis. Shut off the pump once samples have been collected from all sites. Record the total time of release.
  15. Continue recording chloride concentration until stream levels return to pre-release levels. Once measurement in the stream has been terminated, use the probe to measure chloride concentrations of background and plateau samples collected. These samples can then be discarded. Recalibrate the probes, for they may experience electronic drift during the release.
  16. Nutrient Injection Methods: The first nutrient injection (NO3) can be done in conjunction with the conservative tracer injection above. The other nutrient injections (NH4 and PO4) should be done on the next two days, but conservative tracers (Cl or Br) must be added with the nutrients (but the intensive measurement of the conservative tracer concentrations need not be done). On a fourth day do a combined NH4 and PO4 injection if N and P are likely co-limiting.
  17. Mix stock solution of sodium chloride by dissolving 238 g salt per 1 L distilled water. Total volume needed will depend the duration of releases and release rate. Heating the mixture in a water bath aides in dissolution. Mix vigorously and repeatedly for the solution is close to saturation. Make certain the salt is completely dissolved.
  18. Prepare stock solutions of nutrient solutes (sodium or potassium salts). Concentration should be such to produce the desired increase in concentration when mixed in the stream at the upstream station.
  19. Calculate stream flow and necessary release rate to raise stream concentrations appropriately.
  20. Collect a series of background (immediately pre-injection) water samples in mid-stream at several stations over the reach for background nutrient and Cl concentrations. The stations should consist of the upstream reach station (located just downstream from complete mixing of the solute) and several other stations downstream that coincide with stations for the 15N addition. Take 3 replicate background water samples at each station.
  21. Position the Cl probe at the most downstream station to determine when steady state has arrived.
  22. Add nutrient/Cl solution to the carboy and commence injection as described for the conservative tracer injection above.
  23. After plateau (steady state Cl concentrations) has been reached at the downstream site, take at least 5 samples at mid-stream from each station for nutrient and Cl analysis in the same way that the background samples were collected. It is best to do this in separate rounds of sampling, each round consisting of 1 sample from each station from downstream to upstream. This sampling scheme will average over short-term variation in concentrations. The samples should be placed on ice immediately.
  24. After steady state sampling has been completed, the injection can be shut off.
  25. Within 2-3 days, NO3, NH4, and PO4 (soluble reactive P) analysis should be done in the laboratory, if possible. Measurement of the conservative tracer (e.g., Cl) in each sample must also be made, either by wet chemical methods or by ion-specific electrode. We would like to have a measurement precision of + 1 ugN or P/L for these analyses, particularly at sites with low concentrations.

Stable isotopes

  1. Samples of the biota will be collected for determination of stable isotope content in both in streams to which 15N is being added as a tracer.
  2. One transect will be run prior to 15N addition to determine the control del 15N of the various stream components at each station. After the start of the 15N dripper, transects will be run approximately weekly to follow changes in del 15N with time as well as downstream distance. After the dripper has ended, one or two transects will be run to follow the rate of decline of 15N content. Several points on the transect will be above the dripper to be used as control values once the dripper has started. Spacing of stations below the dripper will depend on the size of the stream reach. Stations will be located in riffles unless otherwise specified. In general, try to work from station of lowest expected del 15N to the highest in order to minimize contamination.
  3. Samples representing each of the 15 compartments in the model (NH4, NO3, DON, suspended PON, FBON, CBON, epilithon, filamentous algae, bryophytes, grazers, collectors, filterers, shredders, invertebrate predators, vertebrate predators - the latter two being representatives of trophic level > 3) will be collected. Additional samples may be collected on some dates or at some stations to elaborate some compartments (e.g., CBON split into wood and leaf subcompartments, FBON or suspended PON separated into different size fractions, individuals of other invertebrates from particular functional groups, epilithon on introduced tiles along a transect). Bias your samples toward taxa that are most abundant and can be obtained most reliably. For the standard 15 compartment samples, select one species of grazer, collector, filter feeder, and shredder that can be collected at as many stations and times as possible (you may wish to collect individuals of other species of a particular functional group on some sampling dates/stations if they are also important). Be opportunistic and take many more samples than you can afford to analyze (about 500 per site) because it is impossible to go back and you never know what may turn up in the data later. For example, think of terrestrial critters that depend on your reach and sample them. You can always try to get more sample analysis money later if something exciting appears.
    1. Materials and Supplies
      1. 3 basins, two squirt bottles, scrub brushes (one for reference only), graduated cylinder
      2. turkey baster, 1-mm sieve
      3. Geopump and Gelman filter packs
      4. Small mesh fish net
      5. forceps, filtered lake water
      6. pre-labeled centrifuge tubes and scintillation vials
  4. The following compartments will be sampled on a weekly basis.
    1. Suspended PON
      1. Use Geopump to filter 1 to 4 L of stream water from each station onto a 25 mm diameter GF/F filter fitted in a gelman cassette (need at least 1 mg of ash-free dry mass, so may need to filter larger volumes in streams with low seston concentrations).
      2. Place filter in petri dish. Include one blank filter per sampling period. Dry at 60oC to complete dryness.
    2. Epilithic algae
      1. Using a wire brush, scrub enough rock surfaces to provide at least 1 mg (dry mass) of material from each station. Measure volume of scrubbate and transfer some into 50 mL centrifuge tube. Use a separate, designated brush for the reference station(s).
      2. In laboratory, filter 5mL onto pre-combusted GF/F filter (25 mm diameter), place filter in petri dish and dry (60oC). Include one blank filter per sampling period.
      3. NOTE: It is important to sample the same types of rocks throughout the study reach (to ensure sampling the same type of epilithic community). This will simplify comparison of the 15N content in the epilithic compartment. For example, do not sample "clean" rocks at some stations, and rocks with large amounts of detritus at other stations. If several types of epilithic communities exist (e.g., thin communities in fast flowing sections, thicker communities or detritus-dominated communities in pools or slowly flowing areas), treat them as separate types of samples (epilithon A, epilithon B). Also, consider that certain rocks may turn over or get scoured during high discharge and will have a less mature algal community. Try to sample rocks that are relatively stable.
    3. Bryophytes
      1. Pick clumps of submerged bryophyte species of interest. For bryophytes (as well as other compartments such as filamentous algae below) it is often helpful to have a viewscope to see what you are sampling. Place the collected bryophytes in 50 mL centrifuge tubes with some stream water to keep moist.
      2. Back at the lab, pick off the green tips and place in scintillation vial and dry (60oC). Need about 1 mg of dry material (try to cover bottom of scintillation vial. Also, dry a sample of the entire bryophyte (tips intact).
    4. Filamentous algae
      1. Pick filamentous algae from stream bottom and place in 50 mL centrifuge tube w/ some stream water to keep moist.
      2. Back in the lab, remove as much detritus as possible and place in scintillation vial and dry at 60oC (it might be necessary to collect this material on a GF/F filter first). Need about 1 mg of dry algal material.
    5. FBOM (< 1 mm)
      1. Extract FBOM accumulations that are most available to collector/gatherers from 5 locations at each station using a turkey baster. Pass the material through a 1-mm sieve and collect in a centrifuge tube.
      2. Filter 5ml of the slurry onto pre-combusted GF/F filters, and place in petri dish and dry at 60oC (one blank filter included on each sampling date).
      3. NOTE: Try to avoid collecting inorganic material (e.g., sand) as much as possible (we want the organic rich sediment deposits). On some sampling dates or at some stations you may want to stratify the sampling by habitat type (e.g., pool, riffle).
    6. CBOM (> 1 mm)
      1. Collect by lifting several rocks at each station and catching the "drift" with a small fish net. Dry at 60oC in a scintillation vial. ). Need about 10 mg of dry material.
      2. NOTE: On some sampling dates it would be best to collect separate samples of different types of material if they are important contributors of CBOM (e.g., woody material, leaf detritus ).
    7. Invertebrates
      1. Collect insect samples from rocks using forceps. Sample all common taxa and place at least 5 individuals of each taxa in separate scintillation vials filled with filtered lake water. Allow to sit overnight for gut clearance so that the measured 15N will not reflect unassimilated food particles in the gut. Then dry (60oC) and place in a scintillation vial or in alcohol for later identification. Need about 1 mg of dry material per sample.
      2. NOTE: Concentrate on most common taxon from each functional feeding group which occur throughout the reach in order to get a complete transect (you may wish to collect representatives of a more than one taxon from a particular feeding group on days 7, 21, and 42 if more than one taxon is an important contributor to that feeding group). Also, try to get a complete transect of the dominant organism in each compartment (e.g., grazer, shredder, filterer, predators).
    8. Vertebrates
      1. It is probably best to focus on the youngest age class of salamanders or fish because it is more likely that they will pick up measurable 15N during the study than adults. Depletion may be a problem in many streams, so selection of sampling dates and stations will be important. It might be best to choose one central station to sample from on most dates, and then only do the complete longitudinal survey on the last date (and even then maybe not at all stations). Also, it might be a good idea to discuss with Peterson whether its best to send dissected parts or whole organisms for analysis, given the particular situation at your site.
      2. Place the samples into the drying oven for 24 hrs at 60 C to dry. Once dry, cap and store until shipped to MBL.
  5. 15NH4, 15NO3, 15DON -These samples are collected at most stations on 4 dates (within 12 hours after dripper is started, midway through isotope addition, near end of isotope addition, and within 6 hours after turning off dripper).
    1. Materials and supplies
      1. Ammonium Diffusion Bottles: 4.3-liter widemouth Nalgene bottles
      2. Filter Packs: For NH4 diffusions
      3. MgO Vials: For NH4 diffusions
      4. NaCl and scoop for 200 g aliquots
      5. 4-L jugs for NO3/DON samples
      6. NH4 stock solution
      7. Pipet for 4ml of stock solution
      8. Geopump peristaltic pump
      9. 14.2 cm diameter filter holder
      10. 14.2 cm diameter ashed GFF's
      11. batteries for Geopump
    2. Using Geopump and 14.2 cm ashed GFF's, filter 4 liters water into appropriately labeled Ammonium Diffusion Bottle
    3. Add 200 g ashed NaCl to Diffusion Bottle (use scoop provided to get approx. 200 g)
    4. Cap sample and shake until all salt is dissolved.
    5. Add filter pack to bottle. Note: The filter packs contain acidified GFD's. They trap NH3. Therefore, to avoid contamination, it is important to keep contact with air to a minimum.
    6. Immediately after adding filter pack, add pre-measured vial of MgO.
    7. Immediately after adding MgO, TIGHTLY close diffusion bottle and gently shake to distribute MgO throughout container.
    8. Place bottle in cooler (no ice needed).
    9. Collect an additional 4-liter, filtered sample and place on ice immediately. This sample will be subsampled in Woods Hole for 15NO3 and 15N-DON.
    10. Move to next sampling station and repeat above steps.
    11. NH4 Standard: We will run one standard with each set of ammonium diffusions. The diffusion standard should be started at the same time the ammonium samples are collected. A 4-liter Nalgene bottle containing nanopure water will be shipped with the other supplies. Add 4 mL of the NH4 stock solution (provided) to the 4-liter diffusion bottle then add salt, filter pack, and MgO as detailed above.
    12. Keep the 15NO3 and 15N-DON refrigerated or on ice and send all samples via FedEx to Woods Hole for processing as soon as possible.
    13. IMPORTANT: A separate 200 mL sample should be collected at each location for nutrient analysis (NH4, NO3, and DON). These analyses should be done on site as soon as possible. Precise ammonium values are needed for final 15N-NH4 calculation (but not required before starting NH4 diffusions in the field). Approximate nitrate concentration (if >1 M) is required before processing 15NO3 samples in Woods Hole. If nitrate concentration is < 1 M (14 g/L), precise concentrations are needed before processing samples.

Lysimeter sampling

  1. Method to be added.

Hyporheic sampling

  1. Initial set-up
    1. Materials:
      1. riser (4' to 5' length of ½" PVC tubing)
      2. polypropylene well point
      3. well bottom (5" length of ½" PVC tubing, slotted every ¼" (slots are .006" wide); the polypropylene well point attaches to the lower end of the well bottom, and the riser attaches to the upper end of the well bottom)
      4. silicon stopper (separates riser and well bottoms to prevent exposure of hyporheic water to atmosphere)
      5. 5' to 6' length of 1/8" stainless steel tubing (extends from the well bottom through the stopper to several inches above the top of the riser (top of steel tubing is attached to Swagelok described below))
      6. stoppers (inserted into top of well to prevent rain water from entering well)
      7. 1/8" Swagelok connected to a 1/8" female NPT stainless steel fitting (attached to male NPT below)
      8. 1/8" male NPT stainless steel fitting connected to 1/8" inner diameter plastic hose barb (Luer-lok Cole-Parmer nylon fittings?)
      9. 2" length of 1/8" plastic tubing attached tightly to hose barb fitting
      10. sampling port: 3-way Pharmaseal stopcock (inserted into end of the 2" piece of plastic tubing)
      11. pry bar
    2. Use the pry bar to create narrow gap in streambed cobbles.
    3. As pry bar is removed, insert PVC well as quickly and deeply as possible (30-70 cm).
    4. Deploy wells in a grid consisting of 5-7 rows, 2 to 4 wells per row.
    5. Map well locations and elevations using field level and stadia (referenced to a fixed common point
    6. Allow settling time before sampling.
  2. Hyporheic nutrient sampling
    1. Materials:
      1. 60-ml B.D. syringe
      2. 0.45-m cellulose acetate disposable syringe filter
      3. 60-ml pre-labeled HDPE bottle (for nutrients)
      4. ice
      5. hand pump
      6. vacuum flask
      7. 30-ml pre-labeled glass vial (for D.O.)
      8. parafilm
      9. conductivity and pH meters
      10. Salomet datalogger
      11. MacKereth D.O. probe with stirrer
    2. With the syringe, purge well by slowly drawing at least 30 ml of water through stopcock. Expel this water, and then fill the syringe with 60 ml of water (you should not withdraw more than 100 ml from the well).
    3. Attach syringe filter. Flush 0.45-m syringe filters by expelling 5-10 ml of well water through filter (removes anything attached to filter that might end up in filtrate).
    4. Filter 5-10 ml of sample water into a pre-labeled new or acid washed 60-ml HDPE bottle. Cap the bottle and shake vigorously to rinse. Pour the water out.
    5. Filter rest of the sample in the syringe through syringe filter into the 60-ml bottle.
    6. Keep the nutrient samples cool until returned to camp.
    7. Freeze samples, if they cannot be run immediately.
  3. Sampling of hyporheic dissolved oxygen, pH, and conductivity
    1. With a hand pump, slowly pull water from well into a vacuum flask.
    2. Slowly fill a 30-ml glass vial in the flask and allow to overflow several times into the flask; avoid introducing any air bubbles.
    3. Carefully remove vial from flask (to avoid entraining air bubbles), cap, and seal with parafilm and keep cool.
    4. Measure conductivity and pH on sample remaining in the flask, using portable conductivity and pH meters.
    5. At camp, open vial and measure D.O. with the Solomet datalogger and MacKereth D.O. probe with stirrer; try to measure within 2-3 hours of collection.
    6. Note: small amounts of sediments in samples have not represented a problem for D.O. measurements in the past.

LABORATORY ANALYSES AT TOOLIK

  1. Chlorophyll extraction - River Seston
    1. Materials:
      1. 47 mm GF/C filters
      2. tweezers or forceps to handle filters
      3. graduated cylinder
      4. vacuum filtration apparatus
      5. 15-ml falcon tubes, labeled
      6. styrofoam 15-ml-tube holders
      7. acetone with autopipettor on bottle
      8. cuvettes
      9. cuvette holders, labeled
      10. refrigerator of cooler
      11. foil or cardboard box
    2. Shake and/or invert the 1-L Nalgene bottles to resuspend all materials in the water samples.
    3. Filter 250 ml of water from the 1-L Nalgene bottles through a pre-combusted 47-mm Gelman GF/C filter. Record the volume of water filtered, along with the stream, station, date, and extraction method of the sample, in the chlorophyll book.
    4. With tweezers, carefully place the filters into 15-ml falcon tubes (labeled by river and station).
    5. Place filter for each sample in 15-ml centrifuge tubes with 10 ml acetone
    6. Using the autopipettor, add 10 ml of 90% acetone.
    7. Immediately cover the samples with foil or a box so that they are in total darkness and store in a refrigerator for 24-48 h. Do not freeze or grind the samples, as this will increase the release of chlorophyll.
  2. Chlorophyll extraction - River Scrubs (epilithon)
    1. Materials:
      1. 250 ml plastic bottle (dedicated)
      2. Eppendorf pipettor (at least 5 ml capacity)
      3. 25 mm GF/C filters
      4. tweezers
      5. vacuum filtration apparatus
      6. 15-ml falcon tubes
      7. acetone with autopipettor
      8. refrigerator or cooler
      9. foil or cardboard box
    2. For scrub samples, shake the 56 ml centrifuge tube to uniformly distribute the material. This should be adequate to homogenize most scrub samples. If the scrubs included algal mats, or filamentous algae, use the small blender to homogenize the sample. Blending for 10s should be sufficient.
    3. Pour contents of all five 56 ml cent. tubes into the 250 ml bottle and shake for about 10 seconds.
    4. After shaking to homogenize the sample, withdraw 5 ml using an eppendorf pipette. If there is much material in the sample, then filter only 1 ml (if you had to blend to homogenize, then 1 ml should be sufficient). Filter sample through a 25-mm pre-combusted GF/C filter using the vacuum filtration device. Try to handle filter w/ hand as little as possible to avoid getting any acidity onto the filters
    5. Place filter for each sample in 15-ml centrifuge tubes.
    6. Using the autopipettor, add 10 ml of 90% acetone.
    7. Immediately cover the samples with foil or a box so that they are in total darkness and store in a refrigerator for 24-48 h. Do not freeze or grind the samples.
    8. Conversion from fluorescent units to field units
      1. Use the regression calculated at the start of the field season. The regression uses the 1x fluorometer units only.
      2. The regression gives concentration of Total Chl as ug / L of acetone. This has to be converted to ug / L of water (seston) or ug / cm2 of rock surface (scrubs) as follows:
        1. Seston: (ug Chl a / L of acetone) * (L of acetone used in extraction / L of water filtered)
        2. Scrub: (ug Chl a / L of acetone) * (L of acetone in extraction / L of scrub sample filtered) * (L of water in scrub sample / cm2 of rock area scrubbed).

Ammonium, phosphate, and nitrate - Autoanalyzer

  1. NH4+, PO4-3, and NO3- concentrations are analyzed by the Autoanalyzer RA. If they cannot be done immediately, leave the samples in the refrigerator.
  2. When the analyses are finished, the Autoanalyzer RA will place the bottles back in the refrigerator or in a box outside the trailer. Retrieve those bottles, empty them, add them to the acid bath, and remove them after several hours. Rinse copiously with DI water and set upside-down on a paper towel to dry.
  3. The Autoanalyzer RA will provide a printout or computer file of the nutrient concentration data. Record these data and keep the data printouts or computer files as backup.

Dissolved inorganic carbon (DIC)

  1. Bring the 20-ml teflon septa-capped scintillation vials to the lab for gas chromatography analysis.

Dissolved organic carbon (DOC)

  1. Materials:
    1. clean HCl (a.k.a. Ultrex)
    2. 100-L pipette, tip
  2. Acidify 20-ml water samples to ~ pH 2 with 20 L Ultrex.
  3. Store samples (unrefrigerated) in a sturdy cardboard box. Give to George Kling for analysis.

Alkalinity

(Gran titration method)

  1. Materials:
    1. 50-ml graduated cylinder (or volumetric pipette w/ Brinkman pipettor)
    2. Mettler autotitrator w/accessories
    3. autotitrator calibration standards
    4. wastewater bucket
  2. If you cannot titrate your samples immediately after returning from the field, store them in a refrigerator. The samples should be titrated within a few days (the more alkaline the water, the quicker the sample should be run).
  3. Before sample analysis, bring all water samples to room temperature; pH is strongly influenced by temperature change, so all samples should be the same temperature at the beginning and end of the analyses.
  4. Rinse the Brinkman pipettor with DI and then with approx.10 ml of sample, before dispensing the sample into the special Mettler titration cup.
  5. Analyze 50-ml water samples by Gran Titration on a calibrated Mettler "Auto-Titrator"; record the volume of titrant used and the resulting pH with each iteration of the titration.
  6. Alkalinity is computed by running the macro program which is a part of an excel spreadsheet. Check the output file for correlation coefficient and predicted acid normality) to be sure the data were entered correctly. The correlation coefficient should be greater than 0.98 and the predicted acid normality should be about 0.1N.

Cations

  1. Materials:
    1. clean HCl (a.k.a. Ultrex)
    2. 100-L pipette, tip
  2. Acidify 60-ml water samples to ~ pH 2 with 100 L Ultrex.
  3. Store samples (unrefrigerated) in a sturdy cardboard box.
  4. Ship via Federal Express or hand-carry to MBL, where analysis will be completed.

Anions

  1. Store samples (unrefrigerated) in a sturdy cardboard box.
  2. Ship via Federal Express or hand-carry to MBL, where analysis will be completed.

Epilithic algal scrubs

  1. Chlorophyll (Aquatics trailer)
    1. See Chlorophyll Extraction (sestonic and epilithic) above.
  2. Particulates (Aquatics trailer)
    1. Materials:
      1. 25-mm GF/F filters
      2. tweezers
      3. vacuum filtration set-up
      4. Gelman petri dish
      5. 250 ml plastic bottle (dedicated homogenizer)
    2. Shake each 56-ml scrubbate sample to resuspend all material. Once 5 ml are aliquoted from each of the five 56 ml tubes (for Chlorophyll analysis), combine all tubes into a 250 ml plastic bottle and shake for approx. 10 seconds.
    3. Filter 1 ml from the composite through an ashed 25-mm pre-ashed GF/F filter and the vacuum filtration set-up. Repeat, so that there are two filters for each station (one for particulate nitrogen and carbon, one for particulate phosphorus). Handle the filters with tweezers or forceps, not with your fingers!
    4. Place filter into a labeled Gelman Petri dish and place in the drying oven overnight at 60 degrees C. Dry and store one blank for every 10-15 sample filters.
    5. Store dried filters in plastic bags and ship to MBL.
  3. Algal composites:
    1. materials:
      1. pipette, tips
      2. 20-ml glass scint. vial
      3. Lugol's solution
    2. Shake each 56-ml scrubbate sample to resuspend all material.
    3. Pipette 5 ml from all 5 samples/station (total = 20 ml/station) into a 20 ml glass scint. vial
    4. Add enough Lugol's solution to make a tea colored appearance
    5. Store unrefrigerated in a sturdy cardboard box and ship to MBL.
  4. 15N composites:
    1. Materials:
      1. pipette
      2. 20-ml glass scint. vial
      3. drying oven
      4. 250 ml plastic bottle
      5. vacuum filtration set-up
    2. Shake each 56-ml scrubbate sample to resuspend all material. Once 5 ml are aliquoted from each of the five 56 ml tubes (for Chlorophyll analysis), combine all tubes into a 250 ml plastic bottle and shake for approx. 10 seconds.
    3. Filter 1 ml from the composite through an ashed 25-mm pre-ashed GF/F filter and the vacuum filtration set-up. Handle the filters with tweezers or forceps, not with your fingers!
    4. Place filter into a labeled glass scint. vial and place in the drying oven overnight at 60 degrees C. Dry and store one blank for every 10-15 sample filters. Ship to MBL.

Bioassays of epilithic algae (from Gibeau and Miller 1989)

  1. Analysis of chlorophyll biomass (Aquatics trailer)
    1. Materials:
      1. scrub brush
      2. screw-cap vials (>20 ml)
      3. 90% acetone
      4. fluorometer
    2. Use the incubated discs that were transferred without water. There should be three replicates for each treatment from each station.
    3. Scrub the discs and reserve the scrubbate and the discs for the extraction steps.
    4. Place the discs and the scrubbate in 20 ml of 90% acetone. Place in a dark 4 C refrigerator and extract for 24 h.
    5. Read on the fluorometer in the same manner as described in the chlorophyll analysis section.
  2. Analysis of primary productivity (Aquatics trailer)
    1. Materials:
      1. excess stream water
      2. pipet, tip (?)
      3. 14C-bicarbonate (18 Ci/ml)
      4. scrubber (electric screwdriver fitted with toothbrush head)
      5. 25-mm GF/? filters
      6. tweezers
      7. vacuum filtering setup
      8. light set-up?
    2. Use the incubated discs that were transferred submersed in stream water. There should be three replicates for each treatment from each station. A modified 14C technique is used to assay primary productivity.
    3. Bring the volume of each container up to 100 ml with stream water. Correct the orientation of the disc (i.e., the top should be up, the bottom should be down). (Note: the handling techniques will probably not cause loss of the attached algal mat.)
    4. Inject each sample with 0.2 ml 14C-bicarbonate (18 Ci/ml) and incubate at a constant temperature (10-12 C) and light level (51.7 Einsteins cm-2sec-1 ) for approximately 4 h. ****light set-up? how control light intensity?*****
    5. After incubation, scrub the discs with an electric screwdriver fitted with a toothbrush head. This should remove all algae from the discs.
    6. From each sample, filter (HOW?) a well-mixed 20-ml aliquot of sample through a 25-mm GF/? filter. Handle the filters with tweezers or forceps, not with your fingers! Place each filter into a 10-ml scintillation vial and allow to dry before sealing.
    7. Count the samples on a Packard Tri-Carb 460 C liquid scintillation system utilizing channel radio quench correction. Using the dpm's obtained, calculate primary productivity based on the equations of Wetzel and Likens (1979).

Bacterial production

  1. Preparation
    1. Materials
      1. 15-ml Falcon Centrifuge tubes
      2. test tube rack
      3. 10-ml pipette and bulb
      4. Eppendorf pipette; 1-ml blue tips; 200-l yellow tips
      5. 25-l Hamilton syringe
      6. sterile syringe needles, any thin size
      7. filters - millipore 0.22-m, 25-mm
      8. Millipore filtration manifold, 12-place
      9. vacuum set-up
      10. forceps
      11. 0.2-m sterile disposable filter units
      12. ice, ice bucket
      13. squirt bottle
      14. plastic scintillation vials
    2. Solutions
      1. cold 5% TCA, 200 ml/12 samples ****Make or Buy? ******
      2. filtered (0.2-m) DI water, 1 ml for isotope dilution
      3. filtered (0.2-m) lake water, 100 ml /12 samples, cold
      4. buffered formalin
      5. cellusolve
      6. scintillation cocktail ***** Make or Buy? ******
      7. 14C leucine
        1. stock solution: activity = 314.8 mCi/mmol; concentration = 0.1 mCi/ml=0.32 mol/ml
        2. working solution: 15 l stock solution + 810 l 0.2-m filtered DI water = 825 l working solution
        3. insert thin sterile needle into stock solution rubber cap for pressure release - this needle can be reused
        4. use another sterile needle with 25-l Hamilton syringe for withdrawal add 50 l (30 nM/10 ml sample, 600 ml/12 samples)
  2. Lab Protocol
    1. pipette replicate 10-ml water samples into 15-ml Falcon tubes
    2. add 50 l 14C working solution
    3. Add 200 l formalin to control tubes (kills all bacteria in sample, should get no uptake, hence is control)
    4. Incubate for 1.5 h at lake temp (or 15 C) in dark (preferably in incubator, or if necessary in dishpan with homemade rack placed in lake)
    5. add 200 l formalin to sample tubes (stops bacterial activity)
    6. Pour sample water through 0.2 Millipore filters in filter unit (retains bacteria and can measure 14C uptake)
    7. turn off vacuum
    8. add 5 ml cold 5% TCA
    9. extract 5 (minutes?)
    10. Reapply vacuum.
    11. Rinse filter with 5% TCA
    12. Remove filter unit top
    13. Rinse again with 5 ml 5% TCA (pay particular attention to rinsing edges).
    14. Transfer filter to 20-ml plastic scintillation vial
    15. Add 1 ml cellusolve, let filter dissolve overnight
    16. Add 10 ml scintillation cocktail
    17. Count

YOY grayling

  1. Length and weight analysis
    1. Materials:
      1. 2-phenoxyethanol (0.3 ml per liter of water)
      2. holding tray
      3. digital calipers
      4. paper towels
      5. top loading scale
      6. buckets or 1-L plastic bottles
      7. aerator, aerator tubes and stones
    2. Bring live YOY grayling from the field directly to the lab and transfer them to the holding tray.
    3. Anesthetize using approximately 0.3 ml of 2-phenoxyethanol per liter of water.
    4. Place anesthetized YOY on a paper towel. Measure total length of fish to nearest 0.01 mm with digital calipers (From tip of nose to lower lobe of caudal fin).
    5. Measure wet weight to nearest 0.001 g with top-loading scale.
    6. Hold fish overnight in buckets or 1-L bottles with aerated water.
    7. The following day, release fish in the same place they were caught.
  2. Gut content analysis
    1. Materials:
      1. dissecting microscope
      2. fiber optic light source
      3. scalpel, probe
    2. Under a dissecting microscope, dissect YOY that were preserved in ethanol. Remove stomachs and pick out contents. A fiber optic light under the scope will provide the best results when guts are preserved in ethanol.
  3. Tagging and otolith marking
    1. Materials:
      1. 2-phenoxyethanol (0.3 ml per liter of water)
      2. 26- to 27-gauge needle
      3. 5-cc syringe
      4. ethanol
      5. acrylic paint (various colors)
      6. 200-300 mg tetracycline hydrochloride per liter buffered with sodium bicarbonate (NaHCO3) to approx pH 7
      7. pH paper
    2. On the same day that the fish are caught, anesthetize using approximately 0.3 ml of 2-phenoxyethanol per liter of water.
    3. Tag using subcutaneous injection of acrylic paint; to do this, use a 26-27 gauge needle (sterilized in ethanol) and a 5-cc syringe filled with acrylic paint and inject paint just under the skin on the left, dorsal side of caudal peduncle
    4. To mark the otoliths to determine daily incremental growth rates, place the fish in aerated water containing 200-300 mg tetracycline hydrochloride per liter of water and buffered with sodium bicarbonate (NaHCO3). Hold YOY in this solution overnight to ensure that there are no mortalities due to marking. Warning: This solution can form suds at the surface. YOY can become trapped in these suds and die. To minimize formation of suds, aerate with large tubing that produces large bubbles.
    5. Release fish following day in same location where they were caught; see field methods for recapture procedures.

LABORATORY ANALYSES AT MBL AND OTHER HOME INSTITUTIONS

Particulate organic matter

  1. Materials:
    1. drying oven
    2. dessicator, desiccant
    3. tin disks
    4. two pairs of fine-tipped forceps
    5. aluminum foil
    6. kimwipes
    7. pelletizer
    8. pellet-holding tray
    9. balance (to 0.001 mg)
    10. acetanilide
    11. ashed 25-mm GF/F filters
    12. spatula
    13. Perkin-Elmer CHN analyzer, manual
  2. After transporting the frozen particulate filters back to MBL, place filters in the walk-in freezer until ready to analyze. Make an appointment with Ken Foreman, who will assist with this analysis. Also, use the Manual for the Perkin-Elmer CHN analyzer to assist you; most of the steps outlined below are also discussed in the Manual.
  3. On the day before the filters are to be pelletized, place them in a drying oven at 37C for at least 12 hours. Then transfer the dried filters to a dessicator. Pull a vacuum on the dessicator if possible.
  4. Take the filters to the instrument room (3rd floor, Loeb). Tear off a section of aluminum foil to use as a work surface. Get the tin disks and forceps from the drawers beneath the CHN analyzer and the balance. Clean the forceps thoroughly with kimwipes. Use the forceps to handle the tin disks and the filters; do not use your fingers!
  5. To pelletize a filter, center the filter on a tin disk, sample side up. Make sure you have only one tin disk; they're very thin and it's easy to pick up more than one at once. With the forceps, fold the disk in half so that the filter is on the inside. Fold the disk again lengthwise (you should now have a long thin strip rather than a quarter-circle; see the manual).
  6. Fold the ends of the strip over, and then roll the strip into a sausage or jelly roll shape. If you roll it up too tightly, you may rip the tin, so be careful. When it is rolled up, no part of the filter inside should be visible.
  7. Place the "sausage" (jelly roll if you're vegetarian) into the pelletizing cylinder. Put the cylinder in the widemouth-side of the stage mount. Pull down the handle part way and line up the press with the opening of the cylinder. When they are lined up, pull the handle all the way down and press the sausage.
  8. Lift the handle and remove the cylinder. Flip the stage mount over so that the narrow-mouth side is up. Put the cylinder back on the stage, line up the press and the opening, and pull the handle down. The press will knock the pellet into the narrow-mouth opening.
  9. Using the forceps, remove the pellet and place it into a pellet-holding tray (there are several trays in the instrument room). It is essential that you keep track of which pellet is in which hole in the tray. The holes in the trays are marked (A1, A2, A3, etc.), so use the datasheets provided (titled "AUTO RUN Sample Information (Filters)") to record which pelletized sample is in which hole. Leave the first row (A1-A12) empty; you will put your initial standards and blanks in that row. About every 10 pellets, skip two holes (a standard blank and a filter blank will go in these holes).
  10. Calibrate the balance in the corner, using the instructions provided in the booklet on top of the balance. The calibration weights are in the drawer under the balance. The range setting should be 20 mg.
  11. Pack standard blanks (also called a K factors):
    1. Place an ashed 25-mm GF/F filter on a tin disk, fold them in half with the forceps, and then unfold partially so that there is a crease down the middle of the filter and tin.
    2. Place a counterweight pellet on the left pan of the balance (there are several counterweights in a small plastic box; however, some were made with more than one tin disk and will cause an error on the scale, so keep trying until you get the counterweight with only one tin disk).
    3. Place the creased filter and disk on the right pan of the balance, lower the pan arrest, and zero the balance. Raise the pan arrest.
    4. Using the spatula, place a small amount of acetanilide on the creased filter. Lower the pan arrest and check the weight. Add or remove acetanilide until you have between 2 and 3 mg of acetanilide. Record the weight on a Preliminary Standardization Data Sheet (available in folder on table in the CHN room).
    5. VERY CAREFULLY, using the two pairs of forceps, roll up the disk into a sausage as described above. Make sure you don't spill any of the acetanilide and that you don't rip the tin. Start over if you do either of these things, or else the weight in the next step will not be correct.
    6. Pelletize the sausage and reweigh. If the weight is greater or less than the original weight by more than about 0.010 mg, start over. If not, then record the second weight.
    7. Still using the forceps, place the pellet in a designated hole in the pellet tray. The first five K factor pellets will go on row A (in holes A1, A3, A5, A7, and A9). Subsequent K factor pellets will go within the samples; as you recall, every ten samples or so, you left two holes blank. The first of these holes is for a K factor. So, place the K factor pellet in the first of a pair of empty holes and record the hole number and the weight of the acetanilide in the K factor pellet on the data sheet (use the Preliminary Data Sheet if the pellets are going into row A and the Auto Run Sample Info data sheet if the pellets are being implanted within the samples).
  12. Pack filter blanks:
    1. Using the forceps, place a blank 25-mm GF/F filter that you brought back from the field onto one of the tin disks.
    2. Pelletize the blank filter in the same manner as the sample filters (described above); do not add acetanilide.
    3. Transfer the pelletized blank filter to the pellet tray. Place in one of the empty holes designated for blanks. The first four blanks will go on row A (in holes A2, A4, A6, and A8). Subsequent blanks will go within the samples; as you recall, every ten samples or so, you left two holes blank. The second of these holes is for a blank. So, place the blank pellet in the second of a pair of empty holes and record the hole number on the data sheet (use the Preliminary Data Sheet if the pellets are going into row A and the Auto Run Sample Info data sheet if the pellets are being implanted within the samples).
  13. At this point, you are ready to run your samples. If, however, the CHN machine is already in use or not ready to run, tape your pellet trays shut, label them, and place them in a dessicator with activated dessicant and draw a vacuum on them until the machine is available.
  14. When the machine is available, check with Ken Foreman to make sure that there is enough helium, oxygen, and nitrogen gas, and that the columns are fresh enough to run all of your samples.
  15. loading tray

Total dissolved nitrogen, phosphorus (TDN, TDP)

Total Phosphorus Determination - Acid Persulfate Method – Ian Washbourne (Jan 27, 2003)

Modified by Adrian Green for Arctic Streams analysis – December, 2003

This method is based on previous methods used by Ian Washbourne and Suzanne Thomas, using acid persulfate digestion and analysis on a spectrophotometer. Because our water samples are pre-acidified with HCl in the field (0.1 ml of 6N HCl/50 ml sample), I did not acidify them further. I used acidified DI water, acidified to the same approximate normality as the samples, to make the working standards. I also digested a standard curve with each batch of samples, and ran a non-digested curve each time as well. Changes to Ian’s method are highlighted in bold.

 

Chemicals/reagents                                                                             Amount / Composition. 

Potassium persulfate solution**                                                       < 5 ml (5.0 g / 100 ml, 0.18 M)

HCl(aq)                                                                                                     ~ 2 ml (5.6 N )

A/ Ammonium heptamolybdate tetrahydrate solution*                12.5 ml (2.5 g in 25 ml H2O)

B/ Potassium antimonyl tartrate solution*                                      2 ml (0.5 g in 20 ml H2O)

C/ Sulfuric acid solution                                                                     85 ml (250 ml made up to 1 L with H2O)

Acidified DI for working standards (0.012 N)                              2.14 ml 5.6 N HCl in 1 L DI

 

Working reagent 1**                                                                          100 ml (Add 50 ml of C to 50 ml of 10g / 50ml Ascorbic acid solution.)

Working reagent 2**                                                                          ~ 50 ml (Add 35 ml of C to 12.5 ml of A, mix on the vortex and then add 2 ml of B. Re-mix)  

* stable for months in dark bottle

**make these fresh each time

 

Procedure

-To each vial, sample (10 ml) was added and then concentrated HCl was added in order to acidify to pH 1, (1 drop, 5.6 N) (not necessary for pre-acididfied samples). I only added acid to non-acidified standards and blanks - ACG. The tubes were then vortexed to mix.

-Potassium persulfate reagent (0.25 ml, 0.05g/ml) was then added to each vial and vortexed to mix. Each tube was tightly capped with Teflon lined phenolic caps ready for autoclaving. A few of the tubes were marked with etching or pen in order to create a visual marker to check volume after autoclaving.

-The samples were then autoclaved at 105 °C for 90 minutes. The bottom of the pan was filled with about 2 inches of water before autoclaving. After cooling, each sample was visually checked and taken note of if volume had reduced significantly.   Samples WERE NOT adjusted to account for volume loss, instead if two unchanged replicates of a sample did not remain then the sample was re-run with the next batch.

 Standards

A 10,000mM stock solution was made by adding dried KH2PO4(s)  (1.36g) to a 1 litre volumetric flask, dissolving, adding a few drops of chloroform, and making up to the mark with de-ionised water.

The 50mM stock solution was made by adding 10,000mM (5ml) stock solution to a 1 litre volumetric flask and making up to the mark with de-ionised water.

*Working standards were made with acidified DI (0.012 N HCl) in order to match acidity of samples.

 

 KH2PO4 standard composition.

KH2PO4 Concentration

Volumetric flask used (ml)

Volume of 50mM Stock

0.00 mM

100

0.00 ml

0.05 mM

250

0.25 ml

0.10 mM

250

0.50 ml

0.25 mM

100

1.25 ml

0.50 mM

100

2.50 ml

1.00 mM

100

5.00 ml

1.50 mM

100

7.50 ml

 

 

 

 

 

 

 

 

 

 

 

Organic standard: ATP (C10H14N5O13P3Na2)

- Make a primary 1000µMP standard by dissolving 0.1835g ATP in 1L DI

- Make a working 100µMP standard by dissolving 25 mL 1000µM standard in 250mL DI

(NOTE: but ATP is 7% water, so solution is not 100µM, but actually 93µM)

QCSPEX

Fisher makes a QC standard that contains organic P forms that can be used to check the digestion efficiency of this method.

 

Analysis

-Each standard was transferred to a scint vial (3 replicates, 5 ml in each). To each standard tube working reagent 1 was added (100 ml) with vortexing to mix. After this, working reagent 2 was added to each standard vial, again vortexing to mix. The standards were then left to stand for about 30 minutes, until colour formation was complete, and then re-vortexed and then run on the spec’ at 885 nm to detect for total phosphate concentrations.

 

-Once the standard curve was confirmed to be within acceptable parameters, the samples were reacted with the working reagents in the same way (using 200 mL/10 ml sample of each as opposed to 100 mL to account for the higher volume) and then run on the spec’.

 

-A standard curve was digested along with the samples each time. Two reps were run on all digested samples and standards. Digested check standards and DI blanks were run every 10 samples.

 

Reagents:

Potassium persulfate (K2S2O8) – J. T. Baker 3239-01

Potassium Phosphate (KH2PO4) – Fisher P285-500

Hydrochloric Acid – Trace Metal Grade – Fisher

Sulfuric Acid – Trace Metal Grade – Fisher

 

For analysis on spec:

L-Ascorbic Acid – Fisher A61-100

Potassium Antimony Tartrate C8H4K2O12Sb2*3H20 – J.T. Baker 0864-4

Ammonium Molybdate (NH4)6Mo7O24*4H20 – Fisher A674-500

 

Methods

 ‘Determination of total phosphorus by acid persulphate oxidation’ in Methods of

Seawater Analysis, edited by Grasshoff, Erhardt, and Kremling, 1983

 

‘Dissolved organic phosphorus in the coastal ocean: Reassessment of available methods and seasonal phosphorus profiles from the Eel River Shelf.’ Limnol. Oceanogr., Monaghan and Ruttenberg, 1999.

Cations

  1. Materials:
    1. 60-ml bottle, labeled
    2. graduated cylinder
    3. clean HCl (a.k.a. Ultrex)
    4. 100-L pipette, tip
    5. stock solutions for Na+, K+, Mg+2, and Ca+2
    6. 20 100-ml volumetric flasks
    7. 5-ml volumetric pipette
    8. pipette bulb
    9. 1000-L pipettor (calibrated), several tips
    10. DI water (nanopure for standards)
    11. trace metal grade HCl
    12. 1% lanthanum/5% HCl solution (for calcium)
    13. fume hood
    14. Absorption/Emission spectrophotometer
    15. clean, dry scintillation vials (at least one per sample)
  2. Acidify 60-ml water samples to ~ pH 2 with 100 L Ultrex.
  3. Store for transport and ship to MBL (no refrigeration necessary).
  4. Sign up for use of the Absorption/Emission Spectrophotometer in the Instrument Room (third floor, Loeb).
  5. The week that you plan to run your samples, make standards using stock solutions kept in the back room of the Aquatics Lab, second floor, Ecosystems. For each ion, make standards in concentrations of 100, 200, 300, 400, and 500 parts per million (ppm). Make the standards in 100-ml volumetric flasks (labeled) under the hood. Use trace metal grade HCl, measured in the volumetric pipette. Use nanopure water when making standards.
    1. 100 ppm = 100 l stock, 5 ml HCl, water to fill line.
    2. 200 ppm = 200 l stock, 5 ml HCl, water to fill line.
    3. 300 ppm = 300 l stock, 5 ml HCl, water to fill line.
    4. 400 ppm = 400 l stock, 5 ml HCl, water to fill line.
    5. 500 ppm = 500 l stock, 5 ml HCl, water to fill line.
    6. IMPORTANT NOTE: for each calcium standard, add 1 ml of 1% lanthanum solution before bringing to volume with water. Lanthanum reduces interference with the calcium peak.
  6. When it is time to run the samples on the Absorption/Emission spec, fill out the log book in the Instrument Room (on the little table next to the A/E spec). You will be using the Absorption Lamp and the Continuous Operating current.
  7. Fill the large beaker (inverted on the front of the spec) with DI water (in the carbouy on the shelf to the left of the gas cylinders). Turn on the exhaust fan (switch by the door).
  8. Turn on the air (the knob is on the wall above the gas cylinders). Then, turn on the acetylene. The low stage should stay above 15 (in red), and the high stage should not fall below 70 psi. If a new tank is needed, call Ken Foreman (ext. 507).
  9. Push [Power on]. Do not touch the [Recorder] or [BG correction] knobs. Allow the lamp to warm up for 10 minutes.
  10. Emission methods (use for K+, Na+ analysis)
    1. Adjust the wavelength (with the [Coarse adjust] dial) and slit width for the appropriate ion. For K+, wavelength = 766.5 and slit width = 0.7. For Na+, wavelength = 589.0 and slit width = 0.2.
    2. Push [Flame on]. If an error message appears (due to air in the lines), push [Flame on] again.
    3. Put the sipper tube into the DI in the big beaker.
    4. Adjust the ratio of air/fuel so that it is about 18:40.
    5. Turn the [Signal] knob to EM (emission) and hit [AZ] to auto-zero the spec.
    6. Optimize the wavelength using the high standard from your set of five standards. Place the sipper into the volumetric flask with the high standard. As it is sipping, turn the [Fine adjust] dial slowly back and forth until you establish the maximum readout (you may have to turn the gain up). This is the optimum wavelength.

Anions

  1. Materials:
    1. Dionex chromatographer (at MBL)
    2. stock solutions for Cl-1 and SO4-2
    3. 9 100-ml volumetric flasks
    4. nanopure water
    5. 100- and 1000-l pipettors (calibrated), tips
  2. Make sure that the Dionex is set up to run on the Fast column. If not, contact the lab manager. Also make sure that the Dionex has plenty of regenerant and eluent.
    1. To make regenerant, pour 100 ml of 0.5 N H2SO4 (measure in the dedicated volumetric flask in the lab area adjacent to the Dionex area) into the 2-L volumetric flask dedicated to regenerant (next to the Dionex). Fill to volume with nanopure water and mix well.
    2. To make eluent, weigh 0.4240 g Na2CO3 and 0.0252 g NaHCO3 and pour into the 2-L volumetric flask dedicated to eluent (next to the Dionex). Fill to volume with nanopure water. Mix well (invert and swirl 10-20 times).

Please contact arc_im@mbl.edu with questions, comments, or for technical assistance regarding this web site.