Marsh Ecology Research Program Long-Term Monitoring Procedures Manual by Ducks Unlimited Canada

Marsh Ecology Research Program Long-Term Monitoring Procedures Manual

Delta Waterfowl & Wetlands Research Station Technical Bulletin 2

Marsh Ecology Research Program: Long-term Monitoring Procedures Manual!

Edited by Elaine J. Murkin and Henry R. Murkin

Delta Waterfowl and Wetlands Research Station R.R.1 Portage la Prairie, Manitoba Canada R1N 3A1 Technical Bulletin 2 - 1989

1 Paper No. 54 of the Marsh Ecology Research Program, Delta Waterfowl and Wetlands Research Station and Ducks Unlimited Canada

Preface

Due to the widespread interest generated by the Marsh Ecology Research Program (MERP), the MERP Scientific Team decided to make its monitoring procedures manual available to other investigators. Some of the procedures described herein are new but most have been adopted from other researchers. All have been modified to suit the needs for efficiency, economy, and practicality for long-term, large-scale ecosystem research. We hope tbat some of our lessons, acquired basically by trial and error, will benefit others involved in marsh ecology research. Comments and criticisms of the techniques are welcome. Please direct enquiries to the authors of the individual chapters.

CONTENTS Page Chapter l.

INTRODUCTION TO THE MARSH ECOLOGY RESEARCH PROGRAM Henry R Murldn

Chapter 2.

ANNUAL CUMATIC FACTORS John A. Kodlec

Chapter 3.

PHYSICAL ENVIRONMENT John A. Kodlec

Chapter 4.

HYDROLOGY John A . Kodlec

Chapter 5.

NUTRIENT DYNAMICS John A. Kodlec, Michael P. Stainton and John A . Boughen

Chapter 6.

ALGAL PRIMARY PRODUCTION Sharon E. Gurney and Gordon C. Robinson

Chapter 7.

MACROPHYTEPRODUCTION Arnold G. van der Valk

Chapter 8.

AERIAL PHOTOS AND COVER MAPS Patrick 1 Caldwell and Arnold G. van der Valk

Chapter 9.

DECOMPOSmON Henry R Murldn, Arnold G. van der Valk and Jeffrey W Nelson

Chapter 10.

INVERTEBRATES Lisette C.M. Ross and Henry R Murkin

Chapter 11.

VERTEBRATES William R Clark and Henry R. Murkin

Appendix 1.

FIELD DATA SHEETS

Chapter 1

INTRODUCTION TO THE

MARSH ECOLOGY RESEARCH PROGRAM Henry R. Murkln' Objectives

water levels on the movement and storage of N, P, and C witbin the marsb ecosystem. Tbe more specific researcb objectives are:

In response to the need for long-term multidisciplinary research in freshwater wetlands, the Delta Waterfowl and Wetlands Research Station and Ducks Uulimited Canada embarked on their joint Marsh Ecology Research Program (MERP) in 1979 (Murkin et al. 1984). A Scientific Team from a variety of disciplines was assembled to design and oversee a long-term experiment on the effect of water level manipulations on northern prairie marshes. MERP has three general program objectives:

Hydrology: to estimate tbe terms of the water budget during all pbases of tbe wet -dry cycle: surface water in and out, ground water in and out, change in storage, precipitation, and evapotranspiration. b. Water chemistry: to estimate concentrations of N, P, C, and chloride in surface and interstitial water and to monitor their movement in the water budget throughout the wet -dry cycle. c. Invertebrates: to estimate standing crops of aquatic invertebrates during the wet-dry cycle. d. Macrophytes: (i) to estimate net annual aboveand belowground macropbyte production; (ii) to estimate annual uptake and release of Nand P by living macropbytes. e. Macropbyte litter: (i) to estimate annual production of standing emergent, standing submersed, and detacbed macropbyte litter; (ii) to estimate the annual net loss and/or uptake of Nand P in the standing emergent, standing submersed, and detacbed litter. f. Vertebrates: to monitor waterfowl, blackbird, and muskrat use of wetlands during the wet-dry cycle.

To understand the ecological processes affecting the distribution and abundance of wildlife and plant species in northern prairie marshes; b. To improve practical management of wetlands by providing managers with a better understanding of the structure and function of wetland systems; and c. To encourage students to seek training and careers related to wetland research and management.

The general research objective is to develop new information on the structure and function of northern prairie marshes. Tbese marsbes exhibit cbanges in productivity which correspond to tbe wetdry cycles characteristic of tbe nortbern prairie environment. The wet-dry cycle is tbe fluctuation between drought and flooded conditions resulting from variations in annual precipitation. Tbe cbanges in wetland productivity during this wet-dry cycle bave been described by van der Valk and Davis (1978). Because water regime appears to be tbe dominant factor regulating the productivity of these systems and one which can be practically manipulated by managers, the MERP Scientific Team decided to examine the effects of different

Experimental Design The MERP study area is located on the Delta Marsb in south-central Manitoba. The site consists of 10 contiguous 4-6 ba marsb units (Cells 1-10) created by building a series of dikes along the nortbern edge of the marsh (Fig. 1.1). Besides the diked cells, two undiked areas of similar size within the Delta Marsh (Cells 11 & 12) are monitored as controls. Each diked cell is equipped with a water control structure and electric pump to manipulate

, Delta Waterfowl and Wetlands Researcb Station, R.R. 1, Portage la Prairie, Manitoba, RIN 3A1 1

the schedule of water levels shown in Table. 1. Following a year of baseline data collection (1980), all cells were subjected to a 2-year conditioning period. Conditioning involved flooding tbe marsbes to a depth of 1 m within the cattail (Typha spp.) stands. This flooding was intended to set all cells to the "lake marsh" stage (van der Valk and Davis 1978) to reduce the variability between tbe cells prior to the main experiment. This period was also an unique opportunity to study the marsb response to prolonged above-normal flooding. These levels simulated the natural high water levels that occurred in the Delta Marsh prior to the 1960s. Following drawdown, the cells were reflooded to 3 different levels (Table 2). Under natural conditions northern prairie marshes cycle through varying stages of productivity depending on water levels (van der Valk and Davis 1978). It is hypothesized that the rate of cycle or change in productivity is determined in large by the water depths within the marsh basin. Reflooding the cells to a variety of depths (Table 2) should result in different cycling rates within the study area. Monitoring the movement and storage of nutrients in marshes cycling at different rates will provide important insights into the factors controlling wetland productivity.

Uterature Cited and Suggested Readings MURKIN, H.R., B.DJ. BATT, PJ. CALDWELL, C.B. DAVIS, JA. KADLEC, and A.G. VAN DER VALK. 1984. Perspectives on the Delta Waterfowl Researcb Station - Ducks Unlimited Canada Marsh Ecology Research Program. Trans. N. Am. Wildi. Nat. Res. Conf. 49:253261.

VAN DER VALK, A.G. and C.B. DAVIS. 1978. Tbe role of seed banks in tbe vegetation dynamics of prairie glacial marshes. Ecology 59:322-335. Table 1. Schedule of water levels for MERP experimental cells. Year

Water levels

1980

All 10 cells normal levels of Delta Marsh (247.5Om AMSL)(baseline monitoring of all cells)

1981

8 cells flooded to conditioning level (248.41 m); 2 cells at normal levels of Delta Marsh

1982

10 cells at conditioning level

1983

8 cells drawn down (247.000 m); 2 cells remain at conditioning level

1984

10 cells drawn down

1985 -89

3 cells reflooded to shallow level (247.50 m) 4 cells flooded to medium level (247.80 m); 3 cells flooded to deep level (248.10 m)

Long-term Monitoring Program The Scientific Team has designed a program of standardized methods to monitor the experimental cells throughout the period of study. The basic strategy is to collect a comparable data set from year to year in each experimental cell for each of the major components being monitored. This manual describes the techniques used. Main Transects - Work Lanes To aid in the assignment of sampling stations, each cell is divided into 10 zones. To determine tbe zones, 10 areas of equal width were established between a line 10 m nortb of the soutb dike and a second line 10 m south of the road on tbe nortbern edge of the cell. These zones are permanently marked so they can be located from year to year. Ten transects bave been randomly establisbed within each cell, 1 per zone. These transects or "work lanes" will be used for all types of routine sampling. All sampling will take place as near as possible to tbe work lane. When a sampling point is some distance from the work lane, a patb perpendicular to the work lane will be established.

Table 2. Number of experimental cells within each of proposed water levels following drawdown. Drawdown duration (years) 1 2

Water level aher drawdoWD (AMSL) Sballow Medium Deep (247.50m) (247.80m) (248.lOm)

2 2

, kilomet~ •

NORTH AMERICAN W1LDUFE FOUNDATION

PROPERTY

Fig. 1.1 Location of the MERP Study area on the Delta Marsh

Cbapter 2

ANNUAL CLIMATIC FACTORS Jobn A. Kadlec' 11. Two stations are necessary in order to account for the effect Df the wODded ridge alDng the nDrth side Df the complex. Each of these weather stations includes: i. rain gauge & wet and dry-fall bottle ii. Stevenson screen with max-min thermDmeters iii. class A evapDration pan with max-min thermDmeters

Introduction

Tbe impacts Df aonual vanatlDns in local weatber cDnditiDns on bydrology, pbysical-cbemical cDnditiDns in tbe water and sediments, and many biDlDgical processes are well known. Proximity tD large bodies Df water (e.g., Lake Manit Dba) creates local "lake effects" weatber. Similarly, summer thunderstDrms Dften are very SpDtty in distribution. Consequently, on site measurements Df standard meteorolDgical cbaracteristics are very important. The Dbjectives Dfthe measurements described in this section are: 1) to provide general background data on local weather and 2) to provide data for use in computing water budgets. Because of the importance of hydrologic phenomena to nutrient import and export (see Chap. 5), local measures or estimates of precipitation and evapotranspiration are vital. Precipitation is reasonably well measured by standard rain gauges, which have the added advantage of being widely comparable to other data sets and studies. Evapotranspiration (ET) is less easily estimated. We chose several approaches: 1) standard class A evaporation pans, 2) evaporation pans (class A) installed in the marsh, and 3) estimation of. ET by a modified Penman equation (Eagleson 1970), utilizing data on temperature, wind, solar radiation, and relative humidity. Thus, we installed 2 standard weather stations and 1 special station for wind, radiation and humidity data. Two standard stations were used tD better approximate average weather cDnditions Dver the entire study area. Although most data are recorded daily, only monthly averages are required for integratiDn with mDst other data sets. This had tbe advantage that

IV.

totalizing anemometer

The statiDns are to be mDnitDred daily from 1 May tD 30 October each year (data sheet 1). Rainfall

short term measurement errors tend to average out,

The amount Df rainfall is read directly off the scale (mm) on the graduated cylinder within the rain gauge. Read the level at the bottom Df the meniscus in the cylinder. When there is less than a measurable amount of precipitation (i.e. no complete meniscus is formed) recDrd this as a "trace" by entering "TR" in the field boDk. After the amount Df rainfall has been recorded on the field sheet, discard the cDllected rainwater and replace the graduated cylinder. If there has been a rain event, pour the water collected in the wet-dryfall bottle into a sample bottle. This sample bottle is to be returned tD the lab (see Chap. 5 - Rainfall and Dryfall) for later nutrient analysis. When there is > 25 mm of rain, the cDllected water overflows the graduated cylinder in the rain gauge and collects in the bottDm of the large white container. When this occurs note the amDunt in the graduated cylinder, empty it, and then pDur the water from the white container into the graduated cylinder, nDte the level and add this to the previDus

giving more reliable data Dn the time scale of

amount.

interest.

Maximnm-Mlnimum Air Temperatures Weather Stations Following recording the temperatures Dn the field sheet, reset the thermometers using the magnets supplied.

TwD weather stations are located Dutside the experimental cells. The north station is located nDrth of cell 6 and tbe south station is sDuth of cell

, Department of Fisheries and Wildlife, College of Natural Resources, Utah State University, Logan, Utah 84322-5200 4

Class A Evaporation Pan

ation). This station should be checked daily in conjunction with the other weather stations to ensure the equipment is functiouing properly. Each week the graph paper in each instrument should be changed. On the expired graph record the date and time removed. On each new graph record the date and time installed. On the day the graphs are changed each instrument should be recalibrated according to the appropriate instruction manual. A thermometer and a sling psychrometer will be stored in the instrument shelter for this purpose.

The evaporation pan graduate is used to determine the amount of water added or removed from the evaporation pan. Our present equipment is calibrated in inches (each division is .01 inches). These data will be converted to metric by the computer.

1. Water Added At observation time, if the water level in the pan is below the level of the fixed point within the stilling well, add a measured amount of water to bring the level up to the fIxed point. Record the amount added.

2. Evaporation Pans Within the Marsh Two evaporation pans will be placed in the experimental cells (exact locations will be determined at outset of fIeld season). These pans should be mouitored daily in conjunction with the weather station. Record water added/removed and max/min water temperatures (data sheet 1).

2. Water Removed If precipitation occurs causing the water level in the pan to rise above the fixed point, a measured amount of water should be removed to bring the water level to the fIxed point. Record the amount of water removed.

3. Elevated Totalizing Anemometer An elevated (15 m) totalizing anemometer is located on the west dike of cell 6. Record the reading each day.

3. Net Water Loss The sum of "water added" plus "raint! minus "water removed" equals net water loss. This will be calculated by the computer once all of the com ponents have been entered.

4. Stevens Recorders Two Stevens Recorders, used as a continuous measure of water levels, will be placed near the south end of the borrow ditches in 2 cells. The recorders will be placed in wooden stilling wells. One will be located in cell 6 for the entire season and the other will be moved monthly. The Stevens recorders contain a rotating paper chart that should be monitored in conjunction with the staff gauges (see Chap. 4). Move the recording pen back and forth slightly to create a reference point and record the DATE, TIME and STAFF GAUGE READING. IF IT IS RAINING, DO NOT REMOVE THE LID FROM THE RECORDER. Rain will smear the ink and can ruin several days of data. The recorder should be checked once the rain has stopped.

4. Maximum-Minimum Water Temperatures

Record the maximum-minimum temperatures within the evaporation pans. Reset thermometers. Anemometers

On the totaIizing anemometers, record the current reading. Supplementary Weather Information 1. Weather Station - Cell 6

Literature Cited and Suggested Readings

A third weather station has been established near the control structure on cell 6. This station consists of a hygrothermograph (temperature and humidity) and a recording pyranograph (solar radi-

EAGLESON, P.S. 1970. Dynamic hydrology. McGraw-Hill, New York, NY. 462pp

Chapter 3

PHYSICAL ENVIRONMENT Jobn A. Kadlec'

10 em into the sediments. These temperatures will be entered on the invertebrate data sheet (data sheet 31) along with cell number, transect number, sample site number, vegetation type, water depth, and date.

Introduction Several aspects of the physical environment have important bearings on the measurement or estimation of variables critieal to understanding nutrient cycling. Some of these relate to hydrology, others to sediment structure, and some to chemieal and biological processes or states. Our measures of the physical environment were chosen to provide data to help calculate or inlerpret processes important to nutrient cycling. These include: 1) soil and waler temperatures, 2) water levels, 3) sediment bulk density and organic matter, 4) suspended solids import and export, and 5) bottom contours. Import - export nutrient budgets depend on good hydrologic data (see Chap. 4), an important part of which is the volume of water present at a given water level. Hence data on bottom topography were essential for calculating water level - volume relationships. Similarly, significant amounts of sediment pumped with water into or out of the ponds could have important long term impacts on bottom sediment composition and topography. Finally, sediment bulk density and organic matter determine the relative quantities of water and solids in a volume of sediment, which has an important bearing on the quantities and distributions of many chemical species in the plant rooting volume. Because these physical attributes do not change rapidly with time, measurements are needed less frequently. Input and output of suspended solids in pumped waler (see Chap. 4) were determined monthly, sediment bulk density and organic matter twice a year, and contours once early in the study with later spot checks.

2. Daily maximum and minimum water temperatures will be recorded weekly in representative stands of each cover type (see Chap. 10). 3. Temperatures of interstitial water samples will also be recorded at the groundwater well sites throughout the season (see Chap. 5). Water Levels Water levels in the experimental cells and the main marsh (the south perimeter ditch) will be monitored daily (see Chap. 4 and 5). Cbaracterlzation of tbe Sediments 1.

Particle Size Analysis

An initial survey (Marsh Ecology Research Program - General Soil Survey - April 1979) has provided particle size data. This report is on file at Delta. 2.

Bulk Density and Percent Organic Matter

Soil samples are collected twice a year for determination of bulk density, percent water, and percent organic matter. a.

Sample in the last week of May and September.

Take samples at groundwater (GW) well sites (see Chap. 5).

Use core sampler (to obtain the samples 5 em !D, 15 em length).

Take one fu!! core at each site (if some of core is lost, discard and take another). Transfer the

Soil and Water Temperatures Soil and water temperatures will be monitored in conjunction with the invertebrate monitoring (Chap. 10): 1. Each time an invertebrate station is sampled the following temperatures will be recorded: water column, surface of the sediments (top 1 cm), and

, Department of Fisheries and Wildlife, College of Natural Resources, Utah State University, Logan, Utah 84322-5200 6

core to a wide mouth Mason jar. Label each sample with date and GW well number. e.

In the lab, transfer the sample and aU the water to a pre-weighed container. Record weight of sample plus container and record.

Dry the sample at 1000C to constant weight, cool in a dessicator and re-weigh. Record dry weight (data sheet 2).

Pulverize dry material following drying and take a subsample of about 0.5 g. Place subsample in a pre-weighed combustion pan and record weight of sample plus pan (data sheet 2). Combust in a muffle furnace at 430째C for 24 h, cool in a dessicator and re-weigh. Record fmal weight (data sheet 2).

the sample number on the pan tab. Do not touch the pan or filter; always use metal tongs. Record the sample number and filter-pan weight (data sheet 3). Remove the fUter from the pan and place it on the vacuum fUtering system. Shake the sample and pour 200 ml of sample through the fUter, remove the fUter, and place it back in its original foil pan. Discard the filtrate and remaining sample. Dry the mter and pan for 24 h at 1oo째e. Record the new weight. The difference between the first and second weights on the data sheet is the weight of suspended solids in the sample (will be calculated by computer during data entry). 4. To analyze the sample for the % organic content, combust the above mter in a mume furnace at 430'C for 24 h. Cool in a dessicator and re-weigh. Weight loss = weight of organic matter (OM). wt. of OM/wt. of total suspended solids x 100 = % organic matter.

Determination of Suspended Solids

Bottom Contours

To determine the amount of silt and other suspended solids introduced to or removed from the cells due to pumping or gravity flow through the control structures:

The bottom contours (10 em intervals) of the experimental ceUs have been surveyed by Ducks Unlimited Canada staff. A copy of the contour maps is available in the MERP office. The cells will be resurveyed periodicaUy to monitor changes in bottom contours in response to varying water levels. The staff gauges in each cell were surveyed when they were initiaUy installed near the control structures and in the main marsh. This will allow routine contour checks to be made by using water depths and staff gauge readings. Water depths will be recorded during aU types of sampling in order to allow determination of contour levels. The staff gauges will be adjusted each year to correct any changes caused by frost, settling, etc.

1. During collection of pump samples for nutrient analysis (see Chap. 5) collect duplicate samples at all pump sites during periods of pumping or at aU control structures during periods of active flow. 2. One sample will be used for nutrient analysis (Chap. 5). The other sample should be marked "Silt Analysis" and returned to the Delta lab for analysis of suspended solids. 3. To determine suspended solids; take a pre-dried (100째C for 24 h) fUter and weighing pan and mark

Chapter 4

HYDROLOGY John A. Kadlec' Introduction

The following measurements will be available from the daily monitoring of the weather stations (see Chap. 2): i. rainfall ii. evaporation (class A pan)

Accurate estimates of water budgets are essential pre-requisites for studies of wetland ecosystem nutrient cycles. Hence, the objective of the measurements described below, together with weather and water level - volume data, was to provide water budgets for nutrient budget calculations.

Staff gauges (Fig. 4.1) located near the control structure in each experimental cell and in the south perimeter ditch near the transformer on cell 6 are to be read daily. These are usually done in conjunction with the weather stations each morning. Record the levels on the field sheet marked "Weather and Hydrology Data" (data sheet 1).

The approach to hydrology used was the concept of mass balance: Inputs - Outputs

change in volume

For this approach, all 3 term had to be estimated. Change in volume was calculated from daily records of water level and water level - volume tables (see Chap. 3). Because winds sometimes made water level gauge readings imprecise, 3-point moving averages were used to smooth the data. Inputs of water to diked cells can be in 3 forms: precipitation, water pumped to maintain design levels, and seepage through dike or the sand ridge forming the north end of the cells. Precipitation inputs were calculated from weather records and pumping was metered, providing direct measurements. Seepage was estimated by difference, with checks based on: 1) ground water topography and hydraulic conductivity, and 2) seepage meter spot checks. Outputs of water were evapotranspiration (see Chap. 2 Weather), pumping, gravity flow, and seepage. Gravity flow calculations in 1982 proved inadequate for our purposes and all subsequent water level changes were by pumping. Seepage out was also derived primarily by difference. Because the hydrologic characteristics of different cells in different years were replicated, estimates of the standard error in the seasonal water budgets were possible and proved to be 10% or less.

Pumping

Record the pump meter reading when the water levels (staff gauges) are read. This is usually done in conjunction with weather stations each morning. The time of observation is important and must be recorded. Also note whether the pum p was on or off at the time the meter was read. Record the time and pump meter readings on the field sheet with the staff gauge readings (data sheet 1). The storage disk for this data set is entitled "PUMP METER READINGS". All pumps are set to automatically control the water at design levels. Record the meter reading, time, and staff gauge reading and check for any mechanical malfunctions. 4.

Gravity Flow

During periods of gravity flow to manipulate water levels record the following (data sheet 4):

Water Inputs and Outputs (Water Budget) 1.

Water Level Changes Within The Cells

level at staff gauge immediately before pulling stop logs (see HI below);

date and time stop logs removed and new H' (see below);

date and time stop logs replaced and new H';

Weather Station Data

I Department Fisheries and Wildlife, College of Natural Resources, Utah State University, Logan, Utah 84322-5200 8

(see below)

u. 1Il.

level at staff gauge immediately after replacing stop log;

b. Monitoring Water Storage in the Cells

** •••• *.* •••• _________________ HI

Early in the 1982 field season, a series of wells were installed immediately adjacent to the cells along the north and south borders of each cell. In addition, 1 well was established halfway along the west side of cell 1, and another halfway along the east side of cell 10. Wells to the south, east, and west of the cells were placed along the top of the dike. Placement of the wells north of the cells varies; however, they are as close to the road as the terrain permits. These wells differ from the wells placed within the cells. These wells are made of PVC pipe capped at both ends (Fig. 4.2). Installation consisted of using an auger to drill a hole to the appropriate depth. All wells have the sampling zone completely below the water table, and the sampling zone centered: i. 50 cm. below the groundwater tahle for wells to the south, east and west ii. 30 CDl. below the groundwater table for wells to the north The open tops will extend above the ground surface. At some point during the field season, the elevation of the top of each well will be established (preferably they will be shot in with a transit in conjnnction with DD's survey). The wells should be capped (with PVC end caps or fencepost caps) when not in use. Each well will be labelled with WT (water table well) N, S, E or W (direction from cell), and cell number; ego WTW-l (adjacent well on the west side of cell 1). In 1985, additional groundwater wells were placed in the unflooded portions of cells 3, 7, 9 and 10 to monitor water table topography and all groundwater chemistry. They are similar in design to the adjacent groundwater wells described above, except that the sampling ports are at a single level as in the submersed groundwater wells (see Fig. 4.3). The sampling depth will be approximately 15 cm below the level of the gronndwater (not soil surface). As with the adjacent wells, the elevation of the top of each well will be established each year by surveying. Each well will be designated (and labelled) by WT and location; ego WT 2-9-4, where the numbers are site-cell-zone as with the submersed GW wells. The water table monitoring program will be organized on a seasonal basis, due to the more gradual shift in water table makeup and supply. Sampling will be done monthly for water table level and nutrients in conjunction with the regular groundwater sampling within the cells, and twice

• I I I I I I I

I I I I I I I

• •

• ,. ••••• ------ Lower Water

Surface

Stop Logs HI _ elevation of water surface on high side (staff gauge in cell or south perimeter ditch). H' - elevation of top stop log in structure (determined by measuring from top of control structure - level of top of control structure is marked on each structure). W - width of control opening H' and W will remain constant except when stop logs are added or removed. HI must be recorded at frequent intervals during the period of flow (either by frequent readings of the staff gauges or by continuous recording gauges). The storage disk for this data set is entitled "GRAVITY FLOW READINGS". The format for the computer me is as shown in data sheet 4. The values (HI - H') as well as W will be used to calculate flow rates. F = CW(HI-H') 3/' Additional Physical Measurements 1.

Supplementary Groundwater Sampling

In order to refme estimates of the inputs and outputs for the water budget of the cells, additional physical measurements will be made. These factors will be monitored in order to more accurately defme the water storage of the cells, groundwater levels, and losses through seepage. These additional monitoring programs include: a.

Water table levels Hydraulic conductivity

Supplementary gronndwater sampling adjacent to cells for: i. Nutrients (see Chap. 5) 9

during the season for hydraulic conductivity.

Water Table Levels (Last Monday of each month)

Each month prior to sampling the water table wells (both adjacent to and in the cells) for nutrients, the depth to the groundwater table must be measured (data sheet 5). This measurement is made using a conductivity meter. The measurement is made as follows: i. Use the probe attached to a wood rod with a meter stick on the side. This apparatus will be reserved for this use. u. Plug in the jack from the probe into the

The hydraulic conductivity will be measured at the 22 adjacent water table wells and the WT wells in the dry areas within the cells twice during each field season (late May and early September) (data sheet 6). This sampling should be delayed several days after any water sampling or significant rainfall. The procedure for measuring hydraulic conductivity is: a.

Use the float/plunger apparatus (ie. a dowel calibrated in cm attached to a plastic bottle as a float) designed for this procedure and a stop watch. Note that the vertical doweling is marked at 1 cm intervals.

Measure out 500 ml of water using a graduated cylinder.

Pour the water into the well and immediately insert the float/plunger device. Position your eye at the top of the well. When a cm mark passes the well top, start the stop-watch. Count until 10 cm go by, and stop the stop-watch.

Record the elapsed time and # of cm. If the movement is very slow, record the time to an even cm reading after a period of at least 2 mins. ego 5 COl, 2 min 13 sec.

meter.

Red line (calibrate) the meter. Turn on the meter to the conductivity scale (IX). v. Slowly lower the probe into the well, while watching the meter. Stop the probe as soon as a jump is seen on the meter. Note the reading on the meter stick, then raise the probe and repeat the above process 3 times. All readings should be to the nearest 0.5 COl. VI. Average the reading and record the average. vii. Add 41.1 cm to the reading on the meter stick to fmd true distance (x) from top of well to water surface. viii. Subtract x from the elevation of the top of the well to fmd true level of groundwater in the well.

Hydraulic Conductivity

Ill.

IV.

After obtaining the depth to the water table, completely immerse probe and switch the meter to temperature. Read the field temperature CC) and conductivity and record on the top of the nutrient sample bottle. See instructions for labelling water samples in Chap.5.

A 500 ml water sample should then be collected from the well according to the schedule in Chap. 5.

Literature Cited and Suggested Reading GOSSELINK, J.G. and R.E. TURNER. 1978. The role of hydrology in freshwater wetland ecosystems. Pages 63-78 in Good, R.E., D.F. Whigham, and R.L. Simpson, editors. Freshwater wetlands: ecological processes and management potential. Academic Press, New York, NY. 378pp. KADLEC, JA. 1983. Water budgets for small diked marshes. Water Resources Bulletin 19:223-229. (describes water budget of MERP experimental cells).

There is no computer me for the water table level data set. The data sheets are stored in the MERP office (see data sheet 5).

Fig. 4.1 Staff gauge used to measure water levels within the cells and the main marsh .

. 20

.10

248 . 90

... • ...••

. .••• .•••

248.000

"--

247.790

247.800 247.790

White marks indicate even no. readings

Black marks indicate odd no. readings

247.780 247.7fiJ .70

. 60

247

. 50

...••• ...•• .•••

247.770 247.750

...

•

Fig. 43 Design of water table wells within the cells.

Fig. 4.2 Design of water table wells adjacent to cells. Removeable Cap 1.5 inch ID. PVC Pipe

Snbstrate Surface

Chapter 5

NUTRIENT DYNAMICS John A. Kadlec" Michael P. Stainton', and John A. Boughen " Introduction

Nutrient Inputs and Outputs

In addition to cycling within the wetland, most nutrients are transported in and out of the wetland by water. In this section, we describe the measurements needed to estimate the major waterrelated components of an overall nutrient budget. By multiplying nutrient concentration data by water volume, the mass of nutrients entering, leaving or stored in water can be calculated. Once again, we build on the concept of mass balance:

Inputs - Outputs

Rainfall and Dryfall

Five sample bottles (500 ml) are stored with the weather field sheets. They are labelled: i. Rainwater 1 (north weather station), ii. Rainwater 2 (south weather station), iii. Rainwater 3 (control structure on cell 2), iv. Rainwater 4 (control structure on cell 6), v. Rainwater 5 (control structure on cell 10). If it has rained the previous day, these bottles should be taken into the field when monitoring the weather stations. Collect any water in the wet/dryfall collecting jars at each site (see Chap. 2). Always shake the sam pie vigorously before transferring to the appropriate sample bottle. Do not save the water collected in the rain gauges. Transfer the collected water to the appropriate storage jar in the refrigerator (4째C) in the Delta lab. Rinse the sample bottles with distilled water and return to the drawer with the weather field sheets. If it is the first rain event of the month, it will be necessary to begin 5 new storage jars. Use 2 I plastic bottles and label each with the month and with 'Rainwater 1" through "Rainwater 5". On the first Monday of each month a sample of rainwater from the previous month for each station will be sampled for nutrient analysis. Take the large storage jars from the refrigerator, and subsample by shaking the jar vigorously and then pouring out enough sample to fill 1 of the regular water sample bottles. Label these bottles with the 'RW' prefIX, site (I, 2, etc.), month, and year. Discard the remaining water in the storage jar, wash out the jar with distilled water, and remove the labels. The 5 samples should then be added to the regular water samples taken that day and worked up in the Delta lab (same procedures as surface water

change in storage

Compared to water budgets (see Chap. 4), several differences emerge when determining nutrient budgets: i) water leaving by evapotranspiration does not contain nutrients, ii) nutrients can come from or be lost to solids or biota within the system, and iii) some nutrients enter and leave in gaseous form. Hence we do not expect inputs and outputs via water transport to balance with change in storage in most cases. Within the wetland, nutrients occur in major

pools: surface water, living plants, plant litter, animals, interstitial water, and sediments. The seasonal changes in the pools and the mechanics that control the size of the pools are important factors in the nutrient budgets offreshwater wetlands (Valiela and Teal 1978). All water samples taken by the various procedures described below go through a preliminary series of analyses in the Delta lab and then are shipped for fmal nutrient analysis to the Analytical Chemistry Unit at the Freshwater Institute in Winnipeg, Manitoba.

I Department of Fisheries and Wildlife, College of Natural Resources, Utah State University, Logan, Utah 84322-5200

2 Analytical Chemistry Unit, Canada Department of Fisheries and Oceans, Freshwater Institute, 501 University Crescent, Winnipeg, Manitoba R3T 2N6

Delta Waterfowl and Wetlands Research Station, R.R . 1, Portage la Prairie, Manitoba RIN 3Al 12

sam pies) before shipping to the Freshwater Institute (FWI). There will be no field tern perature recorded

sampling station has been randomly selected within the flooded area on each of the main sampling transects (a total of 10 stations). Beginning in 1986, 5 of these were selected at random for sampling in 1986 and future years. At each sampling station, 3 groundwater wells (Fig. 5.1) have been installed at 3 different depths (15, 30, 45 cm). In the remaining 9 diked cells and cells II and 12, 1 sampling station on 5 of the transects were randomly selected within the flooded area (a total of 5 sampling stations per cell). At each of these stations 1 groundwater well (15 cm depth) was installed. The wells are sampled monthly from 1 May to 30 Oct (Table 1). Groundwater wells may be left in place over winter; however, they should be inspected for damage in early spring each year. To sample a station, take a 500 ml sample from each of the groundwater wells (see Groundwater Well Installation and Sampling Procedures below) and 1 surface water sample (500 ml). To obtain a surface water sample, scoop up a full sample bottle after covering the jar opening with 0.5 mm nylon screening. Be careful not to disturb the bollom before taking the sample. Immediately after each sample is taken, determine sample field temperature by inserting a thermometer directly into the sample bottle. The temperature should be entered on the sample label. Keep water samples in an ice chest or in the refrigerator (4째C), until they are processed in tbe Delta lab. If any wells are broken, they should be removed, the location recorded and a new well put in place before the next sampling period. Repaired wells will be sampled during the last week of each month.

for rainwater samples.

Pumping

During the collection of water samples every Monday (see Table I), collect 1 sample at the pump intake for each cell. Scoop up a bottle of water next to the pump intake. Use the same screen on the bottle as used for surface water samples (see below). Sample whether the pump is running or not. If the pump is not running, turn it on and let it run for 30 seconds before collecting a sample. Label the samples with "PI" or "PO" prefix (see instructions for labelling water samples below), date, and cell number. If the pump is running, take a second sample at the pump discharge. Label these samples with "SUSPENDED SOLIDS", date and cell number. They will be analyzed in the Delta lab (see Chap. 3). 3.

Gravity Flow

Each week during periods of gravity flow, collect a sample (SOO ml) of the water flowing over the stop logs. Wait 10 - 15 minutes after the stop logs have been removed to collect sample. Label sample (use "FI"' or "Fa" prefIX - see instructions for labelling water samples below) and record cell number, date, time, and field temperature (data sheet 4). 4.

Seepage

On the last Monday of each month (see Table I), when the water table level in the adjacent wells is determined (see Chap. 4), collect a SOO ml water sample from the well using a hand pump and a piece of tubing long enough to reach the water table. The procedure is similar to that for wells within the cells (see Groundwater Well Installation and Sampling Procedures below), but varies in that the tubing is not in place at each well and must be carried from well to well. Samples are handled in the same way as the other groundwater samples, and should be labelled with the sample type (WT), the side of the cell (N,S,E,W), cell number, date, field temperature, and conductivity (see instructions for labelling water samples).

Instruction For Labelling Water Samples

Two labels are placed on the lid of each sample bottle: Date: Sample type & location: Well depth (if GW)(cm): Water depth (if SW)(cm): Field water temp ("C):

Nutrient Pools Within The Wetland 1.

Surface and Interstitial Water

Cell 9 has been selected for a detailed survey of surface and interstitial water. Within this cell, 1 13

them 30 cm apart. Always place the well 1 m north of the marker stake. When sampling, always approach from the south side of the stake.

Sample type and location: Example GW2-9-4 b.

To place wells, bore a hole with a 5 cm soil auger to required depth. Carefully push well into hole (avoid tearing mesh) until the plexiglass plate is in contact with the substrate. Run the hoses (air and sample) to the marker stake and attach them with large staples above the water surface. Label the hoses (air and sample) on the stake. If more than 1 well is located at a station, label each hose according to depth of the well and secure all hoses to the same stake. Clamp off the end of each hose. Paint the top of each stake blue and label (in black paint) with "GW", site #, cell, and transect (eg. GW2-9-3).

To take a ground water sample, remove the clamp from the end of both hoses and attach a hand vacuum pump to the sample hose. Adapt a sample bottle cap (Fig. 5.1) so that the sample may be pumped directly into the sample bottle. Fill sample bottle about half way, detach, replace cap, shake and then discard contents. Reattach bottle and ftll completely. Keep water samples in an ice chest or in the refrigerator (4째C) until they are processed in the Delta lab. If no water is obtained by pumping, check the well for broken hoses, plugged tubes, etc. Once sample is obtained, reclamp hose carefully to keep out dirt, invertebrates, etc.

Procedures for Handling Samples In Delta Lab

:.... Transect withln the cell (I - 10) :........ Cell (1 - 12) :............ Sample site withln the cell Cells 1 - 12 except 9 (1 - 5) Cell 9 - (~3,6,8,10 after 1985) Direction from cell (adjacent wells only) (N, S, E, or W) :................. Sample type G W - groundwater SW - surface water PI - pump water into cell (location is indicated by cell # only, well depth and water depth are omitted) PO - pump water out of cell FI - gravity flow water into cell (location is indicated by cell # only, see PI) FO - gravity flow water out of cell RW - rainwater (the only other information on the label will be site 1 to 5, month and year) WT - water table sample b.

Well depth (cm) - For groundwater samples, record the well depth. In all cells except 9, well depth is 15 em. Date - Should be recorded as follows: Month Year ego 2 May 89

The lab must be kept as clean as possible. Small amounts of dirt CaD severely contaminate the samples.

Day

Water depth (cm) - For a surface water samples, measure the water depth immediately above the well.

Field water tempature ("C) Record temperature immediately after sample is taken.

Groundwater Well Installation and Sampling Procedures

Length of well is determined by the depth to be sampled (15, 30 or 45 cm). Where more than 1 well is located at a sampling station, place 14

Transfer the information from the sample labels to the Delta lab sheets (data sheet 7). Sort the bottles (i.e. by sample site within each cell) and keep the sample types (i.e. SW, GW, etc.) separate. Each sample must be assigned a lab number. Check the previous week's lab sheets to determine the next consecutive number. Label each sample with a lab number and Z prefix (eg. Zl28). This number is the code used by the FWI. Record this number on the Delta lab sheet, and fill out the FWI lab sheet (data sheet 8). Use separate sheets for each sample type.

Conductivity and lab temperature: Calibrate the

conductivity meter before use. Submerse probe in sample and move it up and down without touching the sides or bottom of the bottle. Read temperature and conductivity from the meter. Store the probe between samples in distilled water. Change the distilled water every 10 samples. Store the probe between sampling periods in distilled water. c.

pH: Calibrate pH meter with buffer solutions (pH = 7 and 8). Rinse the probe with distilled Submerse probe water between samples. directly into sample bottle. Gently rotate the probe and wait for pH readout to stabilize. Record pH on both Delta and FWI lab sheets. Store the pH probe in distilled water (not buffer solution) between samples. When electrode is not in use, turn the meter off, replace the protective cover over probe making sure cotton plug has been moistened with distilled water. Filtering of samples other than GW: Place the samples to be fIltered in order (by Z#) near the fIltering apparatus. One fIlter is used for C and N analysis and a second for P analysis. Filters are prepared by the FWI and sent to Delta in labelled bottles (ie. C and N, P). Do not touch filters, always handle with forceps, Prior to filtering, label series of small plastic petri dishes (used to store C and N fIlters) with Z#'S. In addition, label 1 dish as a blank. Label a series of small glass vials (used to store P fIlters) with Z #'s (no date necessary) . Again label 1 vial as a blank. Suspended C and N: Place appropriate fIlter on the fIltering apparatus with grided side down. Shake the sample and pour 100 ml into volumetric cylinder. Pour this subsample through the fIlter. Remove fIlter paper with forceps and place in labelled petri dish. Do not discard the filtrate. Once all samples have been filtercd for suspended C and N, place 2 clean C and N filter papers in the "blank" petri dish and handle the same as sample fIlters. Rinse the volumetric cylinder with distilled water between samples. Rinse the fIltering unit between sample types i.e. between SW and GW, etc.

Following fIltering, the fIltrate must be split into 5 subsamples. A variety of coded scintillation vials and a 125 ml bottle are provided by the FWI lab for this purpose. They should all be labelled with the appropriate Z # prior to fIlling. Fill the 4 small vials first and then the 125 ml bottle. If any water is left in the field sample bottle do not discard until all the fIltering and subsampling has been successfully completed.

Filtering GW samples: Suspended C, N and P analyses are not performed on GW samples. They are still fIltered to provide a clean filtrate for further analyses. Do not shake GW samples as they will clog up the filters available in the Delta lab. Prefilter GW samples through the large mesh GF fIlters. Pour small amounts through until the glass filtrate bottle is about half full (250 ml). Change the filter paper if it clogs up. Discard fIlter papers. Refilter using C and N filter papers. Again discard filter papers. The filtrate should then be subsampled as with surface water, etc. Again do not discard sample remaining in the field bottles until filtering and subsampling is completed.

Cleaning equipment: wash all equipment with Decon 75, rinse 2 or 3 times with hot water, and then rinse twice with distilled water.

Preparation of samples for shipment: the C and N petri dishes should be stacked and taped together (with date on tapes). Store in the freezer until shipment to the FWI lab. The vials with the suspended P filters are stored in a wooden box in a cabinet in the Delta lab. When full, it should be sent to FWI lab with the weekly water samples. The small vials should be placed in their appropriate boxes in the shipping cooler. Store the cooler in the cold room (4째C) until immediately before shipment. Place the FWI lab sheets in the cooler with the samples.

Analyses To Be Performed

The following analyses are performed on the samples at the FWI lab (in addition to tern perature, conductivity, and pH measurements taken at Delta):

Suspended P: Place appropriate fIlter paper on the filtering apparatus. Again shake sample, measure out 100 ml and pour through fIlter. Remove fIlter and place in labelled vial. Do not discard the fIltrate . Again at the end of fIltering, place one clean P fIlter in the "blank" vial.

SW, PI, PO, FI, FO, and RW: NH" NO" TDN, SRP, TDP, CI, SO" Susp P, Susp N, Susp C, DIC, and DOC on all samples each month. Na, K, Ca, and Mg on all samples in May and August.

Literature Cited and Suggested Reading

K, Ca, and Mg on all samples in May and August. b. GW and WT: NH" TDN, SRP, TDP, C~ SO" DIC and DOC on samples from all wells (22 adjacent wells and all weUs in cells) in May, August, and October. Same analyses on all samples from all wells within ceUs in June, July and September. N a, K, Ca, and Mg on all samples in May and August.

KADLEC, JA. 1979. Nitrogen and phosphorus dynamics in inland freshwater wetlands. Pages 17-41 in Bookhout, TA., editor. Waterfowl and wetlands an integrated review. Proc. 1977 Symp. NC Sect. Wildl. Soc., Madison, WI. 151pp. Nutrient Dynamics, Part 3. 1978. Pages 155-263 in Good, R.E., D.F. Whigham, and R.L. Simpson, editors. Freshwater wetlands: ecological processes and management potential. Academic Press, New York, NY. 378pp.

Plants and Plant Litter Vegetation and litter nutrient pools wiD be determined in conjunction with the primary productivity (Chap. 7) and decomposition monitoring (Chap. 9).

VALIELA, I. and J.M. TEAL. 1978. Nutrient dynamics: summary and recommendations. Pages 259-263 in Good, R.E., D.F. Whigham, and R.L. Simpson, editors. Freshwater wetlands: ecological processes and management potential. Academic Press, New York, NY. 378pp.

Fig. 5.1 Design of groundwater wells located within the ceUs. Sample outlet (1.5) Air inlet

~~~~~~~~~~~ (I

To hand vacuum pump Sample bottle

~" plexiglass (7x7 em)

Substrate surface

%. plexiglass (30x30 em) l;. 10 plastic tubing P.V.C. pipe (5 em 10) Holes (1 em diameter) Endcap Sample outlet

#-:;;~~;;~~~~~~~: }

Hose clamp Air inlet

Sample depth (15, 30, or 45 em)

- -16

Table 1. Water sampling schedule All cells are sampled on a monthly basis (except adjacent wells that are sampled bimonthly) from 1 May to 30 Oct.

CELLS FIRST MONDAY

NO. OF SAMPLES

9 Pump Water Rain Water

TYPE OF SAMPLES

21 10 5

15 GW, 5 SW, 1 WT 10 PW 5RW

15GW$W,IOPW,5RW,lWT

11 10 12 10 10

5 GW, 5 SW, 1 WT 5GW,5SW 5 GW, 5 SW, 2 WT 5 GW, 5 SW 10PW

-----------------

SECOND MONDAY

10 1 3 4 Pump Water

----------------

THIRD MONDAY

7 8 11 2 Pump Water

FOURTH MONDAY

12 5 6 Pump Water Water table wells adjacent to cells

2OGW,20SW,lOPW,3WT

11 10 10 10 10

5 GW, 5 SW, lWT 5 GW, 5 SW 5GW, 5 SW 5 GW, 5SW 10PW .------------- .--2OGW,20SW,lOPW,IWT

10 10 10 10 22

5 GW,5 SW 5 GW,5 SW 5 GW, 5 SW lOPW 22 GW(May,Aug,Oct) ------------

62 40

37GW,15SW,lOPW (May,Aug,Oct) 15GW,15SW,lOPW (June)uly,Sept)

NOTE: I.

Number of samples may vary slightly due to damaged groundwater wells or other unforeseen circumstances.

ii.

If there is a fifth Monday in a month, sample the adjacent wells that day instead of the fourth Monday. On the fifth Monday of any month also take a set of pump water samples.

iii. On the last Monday of each month sample any wells missed earlier in the month (i.e. wells that were repaired, etc.)

Chapter 6

ALGAL PRIMARY PRODUCfION Sbaron E. Gurney' and Gordon C. Robinson'

during cloudy weather metaphyton may sink well below the water surface.

Introduction

Algae playa critical role in aquatic food chains (Murkin 1989) and in shallow littoral waters the algal contribution to primary production may exceed that of macrophytes (Wetzel 1964). For these reasons, it is essential that we develop a better understanding of algal production in wetland

Objectives The objective of the algal monitoring program is to evaluate the influence of the different water depths on all algal communities within the MERP experimental cells. Within each water level treatment, the effect of stable conditions between 1985-1989 will also be evaluated. The responses will be monitored in terms of:

systems.

Four distinct algal communities contribute to primary production in freshwater marshes. The phytoplankton community consists of freely suspended species within the water column including the tychoplanktors, defined as benthic algal species found within the water column as a result of detachment from their substratum. The tychoplanktors potentially contribute a significant portion of the algal biomass within the water column when samples are collected from within dense beds of macrophytes or during turbulent conditions. In this study no attempt is made to determine the proportion of tychoplankton within the phytoplankton community. Epiphytic algal communities are those species growing attached to aquatic vegetation. The complexity of this community generally increases with colonization time (Hoagland et al. 1982). Initial colonizers are usually adnate to the substrata with colonies of pedunculate forms developing later. Loosely attached filamentous species are often observed late in the colonization proce~ however their direct connection to the macrophyte surface may be minimal. Within aquatic sediments, the epipelic algal community may play an important role in both primary production and in nutrient dynamics. Epipelic species migrate diurnally within the top few centimeters of sediment (Paterson 1986). The mat -forming metaphytic algal

primary production measured through C l • assimilation rates,

b. biomass measured as chlorophyll a (chi a), c.

relationship of water depth, macrophyte density and light attenuation to algal biomass, and

the relative contribution of each algal community to total algal primary production within each water level treatment.

In order to estimate total algal production within each experimental treatment, the following information is required: a.

the relationship between sI?"cific productivity [ILg C fixed • (ILg chi h·I ], light and temperature for each algal community,

the amount of photosynthetically active incident radiation,

light extinction coefficients for sites within each cell, and

estimates of total algal biomass for each community in each cell.

community is de fined as a loose coUection of non-

motile or slightly motile organisms, without any obvious mode of attachment (Round 1981). During extended periods of sunny weather, metaphyton mats can easily be identified near the water surface as they rise by gas bubble accumulation, however,

ar .

Delta Waterfowl and Wetlands Research Station, R.R. 1, Portage la Prairie, Manitoba RIN 3Al

Department of Botany, University of Manitoba, Winnipeg, Manitoba R3T 2N2 18

Sample Site Selection 1.

Phytoplankton

In each of the 10 diked cells, light extinction measurements are conducted on a 3-week cycle on 2 randomly selected lanes. At 10 m intervals along each lane, a LI-COR solar monitor (LI-1776) with an underwater qnantum sensor (LI-19OSB) is used to measure the amount of photosynthetically active radiation above the vegetation, at the water surface, and at 5 cm intervals within the water column to a 25 cm depth (data sheet 9). Continuous hourly monitoring of incoming photosynthetically active radiation is recorded at the weather station at the south end of cell 6 (see Chap. 2) with a LI-COR solar monitor (LI-1776) equipped with a quantum sensor (4-19OSB). Twice weekly, the data are recorded (data sheet 10), the sensor cleaned, and battery levels checked.

Within each of the 10 diked cells, 3 sampling sites are randomly chosen; 1 site is located in near maximum water depth, the second site at a shallow depth (approximately 28 cm) and a third at an intermediate depth. These sites are permanent stations used for a 3-week interval sampling schedule.

Epiphytic Periphyton

Site selection is stratified by water depth and cover type. Within each of the 10 diked cells, sites are selected to cover all major vegetation types and the entire water depth range. Epiphytic sampling sites are located adjacent to the macrophyte permanent quadrats (see Chap. 7). Artificial substrata made from 0.6 cm diam. extruded acrylic (Goldsborough et al. 1986) are used for the collection of epiphytic algae. At the beginning of each field season, acrylic rods (prescored at 3 cm intervals to allow subsampling at specific depths) are placed at the sites to allow a 3-week colonization period before the first sampling date. Forty rods (5 rods x 8 sample periods) are pushed vertically into the sediment. Rods are equally distributed amongst the vegetation within the site (1 m') to simulate light conditions available to epiphytic algae growing on

light measurements above the emergent vegetation, at the water surface and at 5 em intervals through the water column. The readings are made on the north, south, east, and west sides of the sampling site,

b. Above-water vegetation density is estimated using a seale of 0-5, with 0 representing open water and 5 dense emergent vegetation where little light penetrates the water surface,

Epipelic Periphyton

For those cells containing non-vegetated submerged sediments, 1 site per cell is randomly selected for epipelon sampling. The total area of non-vegetated submerged sediments within a cell is dynamic, therefore the sampling sites are not permanent throughout the field season. Fluctuations in density of submersed macrophytes and metaphytic mats necessitate estimating the area of nonvegetated submerged sediments at regular intervals. 4.

All Sites

At all algal sampling sites the following parameters are recorded on each sampling date (data sheets 11-14):

natural substrata.

Light Measurements

above-water vegetation type using a % cover value. Live and dead stems are differentiated in this estimate,

density of submerged aquatic vegetation using an index of 0, low, medium, or high based on the % of the water column supporting submerged aquatic vegetation: 0 = 0%, L = 133%, M = 34-66%, H = 67-100%,

water depth averaged over the sampling site,

water temperature, and

% coverage of metaphytic mats within the sample site.

Phytoplankton

Metaphyton

Due to the temporal variability of metaphytic mat development, estimates of metaphytic cover are made on a 3-week cycle. The % cover is estimated at 10 m intervals along each work lane in each cell. From all sites supporting metaphytic mats, 3 sites are randomly selected per cell for biomass estimates. Field Sampling Methods

At each sampling site, a plexiglass tube (5.8 cm 19

10) is used to collect an integrated water sample

to collect submerged macrophytes from the water column. Plants are collected from an area which has a water depth and vegetation density similar to that present in the permanent quadrat, although not within 5 m of the permanent sampling quadrat sites. The clipped vegetation is labelled and returned to the laboratory for surface area measurements (data sheet 13). Surface areas are calculated using formulae for the nearest geometric solid. These surface area determinations and the stem densi?, data are used to calculate the surface area per m . The total submerged surface area within each cell is calculated using data collected from all sampled quadrats.

(data sheet 11). Care should be employed to minimize the amount of disturbance to the aquatic vegetation so as to reduce the amount of tychoplankton in the sample. For estimates of chi a, 4 500 ml subsamples are taken from the integrated sample. An additional 500 ml subsample is taken when conducting carbon assimilation experiments (see below). Tbe 30 phytoplankton sites are sampled over a 3-day period. Water samples are collected in the morning and ftltered onto 3.3 cm GF IC ftlters and frozen that afternoon to minimize the breakdown of chi Q. 4.

Epiphytic Periphyton 5.

At3-week intervals, 4 replicate acrylic substrata, collected from the north, soulb, east, and west sides of the site, are subsampled for algal biomass. The number of sUbsamples collected from each rod is dependent on the water depth (data sheet 12). For each of the following water deptb ranges, the following 3-cm segments from each of the 4 replicate rods are sampled: i. 1-14 em: collect 1 segment from the middle of this rod, II. 15-24 cm: collect 2 segments, 1 from a 3 cm depth and 1 segment from the bottom of the rod, Ul. 25-34 em: collect 3 segments, 1 from a 3 cm depth, 1 from the middle of the rod, and 1 from tbe bottom of the range, and iv. > 34 cm: collect 4 segments, 1 from a 3 cm depth, one from the bottom of the rod, and 2 segments which are equally spaced between the top and bottom segments.

Epipelic Periphyton

On a 3-week cycle, area estimates of unvegetated submerged sediments are made for each cell. These estimates are conducted concurrently with the estimation of metaphytic cover (data sheet 14). For biomass determinations, 3 replicate sediment samples are collected at each site, using a plexiglass corer (5.8 em ID) (data sheet 11). An aspirator is employed to remove the top few cm of the core to a 500 ml sample bottle. Samples are then returned to the laboratory and settled in darkened beakers (10 cm ID). The following evening (when the algae are not migrating to the sediment surface), water overlying the sediment is carefully removed so as not to disturb the sediment, 10 em diameter lens tissue traps are placed over the sediment surface, and the beakers are returned to natural light conditions under a plexiglass cover. Because epipelon migrate only to the sediment surface and not into the water column, care must be taken to ensure that sediment samples are damp but not flooded. This may involve the removal of overlying water throughout the trap sampling period or addition of water under very warm conditions. Due to temporal variability in migration times of individual species, tissue traps are removed and replaced with new traps at 10:00 am, 2:00 pm and 7:00 pm on the day following placement of traps. When conducting biomass and productivity experiments simultaneously, traps are placed in 100 ml offtltered site water (as described above), shaken vigorously, then dispensed into culture tubes for assimilation experiments, with a portion (usually 25 ml) filtered through a GFIC filter, frozen and used for chi Q analysis. When C- assimilation experiments are not being conducted, tissue traps are placed directly into 14.8 mllabelled specimen bottles and frozen for biomass determinations.

Needle nose pliers are used to break the segments at the score mark. Biomass samples are placed individually into 14.8 ml specimen bottles labelled with the site number, depth level and replicate number. Samples collected for C1â&#x20AC;˘ assimilation experiments (see below) are placed in culture tubes containing 25 ml of ftltered site water. Site water is collected 1 day prior to rod sampling, filtered through 0.45 /Lm membrane filters and stored at 5Â°C in the dark until required. For whole cell production estimates, the surface area of available substratum for epipbytic growth must be determined. At the end of June and the end of August, the stem density data from the permanent quadrats (Chap. 7) is used to determine what species are present in those quadrats adjacent to the epiphyte sites. For each emergent species, 2030 stems are clipped at the water surface and again at the sediment surface. A 0.25 m' quadrat is used 20

Metaphyton

transfer of 4 ml to a clean cuvette. The ftlter (MSI glass fiber, 0.8 /lom pore size) should be changed after each set of replicates. Expel residual extract from filtering device between ftltrations.

At 10 m intervals along each lane within the diked cells, % cover of metaphyton is estimated by placing 4 0.25 m' quadrats on either side of the transect (Gurney and Robinson 1988). Water depth and emergent and submerged macrophyte densities are also recorded at each site (data sheet 14). Following the metaphyton cover estimates, biomass sampling is conducted at the 3 biomass sites in each cell (data sheet 11). Four replicate biomass samples are collected at each site to determine the relationship between % cover and biomass (measured as chl a). At each site, a small (ca. 50x30x4cm) piece of styrofoam is submerged and used to bring the metaphytic mat above the water surface. A copper cylinder is used to remove a 1.54 cm' area of the mat. For biomass estimates, 1 core is placed into each of 4 14.8 ml specimen bottles. When productivity experiments are being conducted, cores are placed individually into culture tubes containing 25 ml of fIltered site water. In addition to the standard list of parameters recorded, mat depth is determined. This measurement is made from the water surface to the top and then bottom of the mat. An additional light reading is taken under the mat when conducting light extinction

Clean sides of cuvettes, then read absorbance of samples at 665 om and 750 nm in a spectrophotometer (data sheet 15).

Add 100 /lol of acid stock (to make a 500 ml stock solution: add 2.0 ml of concentrated HCI to 498 ml of distilled water, then fIlter stock solution through a 0.45 /lom membrane filter), invert cuvettes 2-3 times to mix the acid in the cuvette, cover samples and allow samples to acidify for 1 h.

Read absorbance again at 665 and 750 om, making sure the sides of the cuvettes are clean.

Concentrations of chl a are calculated as described in Marker et al.(1980).

Notes: -Keep a lid over cuvettes to reduce evaporation and light contact with samples -If samples are very concentrated, you may have to dilute your samples with filtered methanol then correct your absorbance reading accordingly. -Do not re-use scratched or discolored cuvettes. -If the 750 om reading after acidification increases substantially ie. 2-5 times, it is possible that you may have organic acids precipitating in your samples. To correct this, refilter the contents of the cuvette, place ftltrate in new cuvette and reread the absorbance at 665 and 750 nm. Your values will drop if you did have a precipitate forming. -Your readings at 665 om should drop after acidification unless you had no intact chI a in your sample (rare), your 750 values may go down or up a small amount. If in doubt, rerun the sample using another 4 ml of extract.

Primary Productivity Experiments

measurements.

Sample Storage

All algal samples must be kept cool and stored in the dark until they can be transported back to the laboratory. Samples collected for chl a analysis, must be placed in the freezer immediately upon return from the field. Samples to be used for productivity experiments should be placed in the dark at 5째C until they are inoculated with isotope. Laboratory Procedures 1.

ChI a Analysis

Add 90% methanol to algal samples (8 ml per 500 ml phytoplankton sample; 8 ml per 3 em segment of acrylic rod; 8 ml per 22.8 cm' epipelon sample area; 20 ml per 1.54 cm' metaphyton core), then shake gently (GF/C's) or vortex (rods, epipelon and metaphyton) 3 times over a 24 h period keeping samples in the dark at approximately SOc.

a. Algal samples are inoculated with a standardized amount of C 14 bicarbonate and placed in a water filled incubator for 2 h at ambient water temperature.

Using a sterilizing syringe system with a 13 mm filter, prepare a blank cuvette (1Ox10x45 mm polystyrene) with 4 ml of 90% methanol. Then, filter enough of each chlorophyll sample to allow

Replication of a range of light intensities is achieved by submersing samples at different distances from a high pressure sodium lamp,

generating pbotosyntbeticaU1 active radiation (P A.R.) up to 1900 ",Em·'s. ALI-COR (LI1n6) solar monitor with an underwater quantum sensor (LI-I90SB) is used to measure the light intensity for each replicate row in the incubator. c.

assemblages, with emphasis on the diatoms (Bacillariopbyceae). Am. J. Bot. 69:188-213. GOLDSBOROUGH, L.G., G.C. ROBINSON, and S.E. GURNEY . 1986. An enclosure/substratum system for in situ Arch. ecological studies of periphyton. Hydrobiol. 106:373-393.

Samples are then removed from the incubator and flltered through a 0.45 ILm membrane filter and rinsed with distilled water.

Filters and algae are fumed over concentrated HCI for 1 min to remove residual inorganic C" and placed in vials containing scintillation fluor.

A Beckman LS3801scintillation counter, with HNumber correcting quenching, is used to

GURNEY, S.E., and G.C. ROBINSON. 1988. The influence of water level manipulation on metaphyton production in a temperate freshwater marsh. Verh. Internal. Verein. Limnol. 23:1032-1040. MARKER, A.F.H., EA. NUSCH, H. RAI, and B. RIEMANN. 1980. The measurement of photosynthetic pigments in freshwaters and standardization of methods: conclusions and recommendations. Arch. Hydrobiol. Beih. Ergebn. Limnol. 14:91-106.

determine sample radioactivity.

C-assimilation rates are then calculated using the formula (data sheet 16): ILg C fixed' cm" or 1'1 • h· 1

DPM(S) x C x 1.05 DPM(T) x M x T

MURKIN, H.R. 1989. The basis for food chains in prairie wetlands. Pages 316-338 in van der Valk, A.G., editor. Northern Prairie Wetlands.

Where: DPM(S) specific radioactivity of sample, corrected for dark uptake. DPM(T) = specific radioactivity added to the samples. C = total inorganic carbon in incubation water derived from alkalinity (APHA 1980), pH and temperature (data sheet 17). 1.05 = CI • discrimination factor. M = area (cm') or volume (I) of algal sample. T = incubation time (h).

Iowa State University Press, Ames, Iowa.

PATERSON, D.M. 1986. The migratory behavior of diatom assemblages in a laboratory tidal micro-ecosystem examined by low temperature

scanning electron microscopy. Diatom Research 1:227-239.

PLATT, T., C.L. GALLEGOS, and W.G. HARRISON. 1980. Photoinhibition of photosynthesis in natural assemblages of marine phytoplankton. J. Marine Res. 38:687-701.

Specific productivities IILg C fixed' (ILg Chi h·ll are then calculated as a means' of normalizing data (data sheet 18). The standardized data are then used to develop photosynthetic/irradiance (P vs I) SAS non-linear curves for each community. regression will be used, where possible, to describe the parameters of the P vs I curves using the model of Platt et al. (1980).

ROUND, F.E. 1981. The ecology of the algae. Cambridge University Press, London, UK, 653pp.

I •

WETZEL, R.G. 1964. A comparative study of the primary production of higher aquatic plants, periphyton, and phytoplankton in a large, shallow lake. Intern. Rev. ges. Hydrobiol. 49:164.

Literature Cited and Suggested Reading AMERICAN PUBLIC HEALTH ASSOCIATION (APHA). 1980. Standard methods for the examination of water and wastewater. 15th ed., Washington, D.C. 1134pp. HOAGI.AND, K.D., S.c. ROEMER, and J.R. ROSOWSKI. 1982. Colonization and community structure of two periphyton 22

Chapter 7

MACROPHYTE PRODUCfION Arnold van der Yalk' Introduction

belowground biomass begins to increase quickly after the minimum has been reached.

Determination of aquatic macrophyte annual net primary production is vital to the understanding of the dynamics of freshwater marshes. Macrophyte biomass, both live and dead, is a major storage compartment for carbon, nitrogen and phosphorus in a marsh and a major potential energy and nutrient source for the faunal component of the marsh ecosystem. Macrophyte communities are also essential structural components of the habitat of both invertebrates and vertebrates. The major objective of our long-term monitoring of aquatic macrophytes is to determine the impact of the wet-dry cycle on macrophyte above and belowground net annual production. Standard harvest techniques are used beeause they are the most direct, simple and reliable techniques available for estimating net annual primary production of macrophytes per unit area.

.b.

Fall Sampling Period Last 2 weeks of September and first week of October (begin September 18). Cattails should be visibly senescing, i.e. turning yellow, at the time this sampling period begins.

Sample Site Selection Randomly select 2 sample sites along each transect (worklane) in each cell before beginning the spring sampling. To select the sample sites on a transect (workIane), pick 2 random numbers between 0 and 100 (data sheet 19). Each random number represents the position of the sample site in percentage of the total lane length (from the eastern boundary of a cell). For example, if the random number is 72 then the sample site will be 72% of the distance along that transect from the eastern boundary. (Remember there is a 10 m buffer zone around the edge of each cell). On the detailed topographic map of a cell, locate the worklanes and determine their length. Multiply the length of a worklane by the 2 random numbers for that worklane expressed as a decimal Craction; e.g. if the worklane is 150 m long and the random numbers are 16 and 72, then the 2 sample points will be located along that transect at (150 x 0.16) or 24 m and (150 x 0.72) or 108 m from the eastern boundary of the transect (or 34 and 118 m Crom the edge oC the east dike, respectively). In the field these sampling points should oe located by pacing oCC the appropriate distances along the transect. Altogether there should be 240 belowground macrophyte samples collected in the spring and fall, i.e. (2 samples/transect)*(lO transects)*(12 cells).

Sample Site Marker Stakes Place a marker stake in the spring at each sample site. These same sites will be used in the fall. On the stake record the Collowing: MBG (indieates macrophyte belowground site), site number, cell number, and transect number.

Belowground Macrophyte Production An estimate of net annual belowground macrophyte production requires that samples be collected twice during the growing season: initially in the late spring when shoot initiation has used up most of the belowground store of carbohydrates (i.e. at the time of minimum belowground standing crop) and again in the fall when underground biomass has reached its seasonal maximum (van der Yalk and Davis 1978a). The difference between fall and spring standing crops provides an estimate of the net belowground production. I.

Field Sampling Methods

Spring Sampling Period To ensure that spring belowground samples are taken at the time that underground reserves are reduced to a minimum, spring sampling begins when the eattails (Typha spp.) start to flower, the time when underground reserves are lowest for this species. Spring belowground samples should be collected as quickly as possible, i.e. in less than 2 weeks, beeause

Department of Botany, Bessey Hall, Iowa State University, Ames, Iowa 50011 23

Within a cell the sample site numbers run from 1 to 20. Sites are numbered starting at the northeast corner ofthe marsh with sites 1 and 2 along transect 1, sites 3 and 4 along transect 2, etc. Along a transect the eastern-most site always has the lowest number. Yellow flagging should be attached to each stake. If a MBG sampling site falls at the same location as a permanent quadrat, the MBG sampling site should be moved 5 m west. e.

Coring Sites All spring and fall cores will be collected from a 10 m transect running due north of the sampling stake. Coring sites will be selected randomly at 1 m intervals along this transect. Three cores will be collected in the spring and 3 cores in the fall. In the spring select 3 random numbers between 1 and 10 at each sampling site and collect cores at these 3 positions along the coring transect (data sheet 20). In the fall collect cores at 3 additional positions.

Sampling Schedule During a sampling period do not collect all the MBG samples in a cell at once. Transects in each cell are grouped into 5 pairs (1 and 2, 3 and 4, 5 and 6, 7 and 8, and 9 and 10) for a total of 60 transect pairs. Each transect pair should be assigned a random number between 1 and 60 and the transect pairs sampled in that order (data sheet 21). This schedule of sampling avoids introducing a seasonal bias into the samples which would be present if all sam pies were collected systematically by cell.

the coring transect at the site. Also record average water depth at the coring sites. Information from the field book should be transcribed each day to laboratory data sheets (data sheet 22). 2.

Root Washing and Sorting

On return to the Station, MBG samples should be placed in the shade near the root washer if they are to be washed immediately or in the cold room (4째C) if they are to be cleaned later. The root washer is simply a rotating cylinder in a tank with a flow-through water system. The cylinder has 4 compartments which each holding a sample. Each compartment has a screen door which allows dirt to be washed from the sample as the cylinder rotates. Before loading cores from a bag into the washer, remove the label from the bag and place it in the clip opposite the compartment in which the sample is to be placed. If, after washing in the root washer, the sample is still not free of all sediment, it must be washed manually using the high pressure sprayer. If washed samples are not to be sorted immediately, the plant material is placed in a plastic bag with the original label inside. These cleaned samples are then stored in the cold room (4째C) until they can be sorted.

b. All samples are sorted into live and dead components. Reference samples to aid in identifying roots and rhizomes are available for all the common species in the cells. Persons working on sorting should familiarize themselves with these reference samples. Any uncertainties ahout whether a root or rhizome is live or dead, should be resolved with the person in charge of MBG sampling. After sorting has been completed, live and dead plant material are placed in separate paper bags and labelled with: MBG, sample site, cell, transect, live or dead, and date. Each sample is then dried to constant weight at BOoC in the drying room and weighed to the nearest 0.01 g. All samples are retained for grinding.

Coring As described above, each sample consists of 3 cores. These cores are 15 cm in diameter and taken to a depth of at least 20 cm. At most sites 20 cm is deep enough to sample the entire root and rhizome zone. However, as samples are collected in the field, cores should be checked to make sure that the whole rooting zone is being sampled, i.e. cores may need to go deeper than 20 cm at some sites. Clip all live and dead stems off at ground level (this can be done before or after the core is taken, but it is usually easier to do it before). The 3 cores collected at each site are placed together in a heavy-duty plastic bag. Apply strips of masking tape near the open end and label with MBG, sample site (1 to 20), cell number (1 to 12), transect (1 to 10), and date (day, month, year). In a field book record the information on each label plus all plant species present along

Grinding Samples In preparation for nutrient analysis, all sam pies are ground in the Wiley mill. The entire sample is ground and placed in a Whirlpak bag. If the sample is very large, take a subsample (about 20 gms) and discard the remaining sample. Ground samples are

numbered consecutively and are given a prefix indicating the type of sample, i.e. 42MBG89 for tbe 42nd sample ground in 1989. Ground samples from a given sampling season are stored togetber in boxes appropriately marked, e.g. MBG Spring 1989.

in each cell are grouped into 5 pairs (1 and 2, 3 and 4, 5 and 6, 7 and 8, and 9 and 10) for a total of 60 transect groups. Use the same schedule as belowground macrophyte samples (data sheet 21) to avoid introducing a seasonal bias into the samples.

Aboveground Macropbyte Production

Maximum standing crop is used to estimate annual aboveground macrophyte production. Because most prairie emergents reach their maximum standing crop in late July or early August, sam piing will take place during this period. A single clip-plot method underestimates net primary production, however this estimate can be corrected using the data from the turnover plots (see below). 1.

Clip Plots

Sam piing period Samples are collected around tbe last week of July (July 18). Emergents like Typha should appear fully grown before this sampling is begun.

Sample sites Pick 5 random sites (only 4 will normally be used) per transect using the same technique as used for belowground macrophyte sample site location (see data sheet 19, use 5 sample sites instead of 2). Do not use tbe sampling points selected for the belowground macropbyte samples. These 2 sets of samples are completely independent of each other. Locate the fIrst 4 random sample sites selected along each transect in tbe fIeld. If a sample site falls on tbe site of a belowground macrophyte sample or permanent quadrat, use tbe 5th random sample site as an alternate sampling site. Altogether 480 quadrats will be clipped each summer, i.e. (12 cells)*(4 samples/transect)*(10 transects/cell). Samples are numbered in a cell starting with the northern most transect and along a transect are numbered from east to west, i.e. transect #1, samples I, 2, 3, and 4.

Dry Sites: clip all standing vegetation (living and dead) within the quadrat. Only plants rooted within the quadrat are included, not plants hanging or falling over the quadrat. Large hand clippers are used to cut the plants by placing the open blades firmly on the substrate and tben closing the clippers. It is important tbat all quadrats be clipped as close as possible to the substrate and that this be done in a uniform manner. Following clipping, the vegetation should be separated by species or groups of species (as described below). The following emergent species are always separated; Scholochloa

festucacea, Typha spp., Phragmites australis, Scirpus lacustris, and Carex atherodes. Any other species with a cover of > 25% in a plot should also be separated by species. All species with a cover of < 25% in a quadrat are combined into 1 group and labelled, "MINOR SPECIES". All 6 major species, any other dominant species, and the "MINOR SPECIES" are also separated into live and dead fractions. Live and dead material of each species should be placed in separate bags and labelled with MAG (macrophyte aboveground), sample site

number, cell number, transect number, species name, live or dead, and date. In the field book record all subsamples at the site. Paper bags may be used for small samples, but burlap bags, tied shut with twine should be used for large samples. Cardboard labels sbould be tied to the burlap bags.

ii.

Quadrat Location

A I x I m (Typha, Phragmites, Scirpus) or a 1/4 x 1/4 m quadrat (all other species) is located at each site with its northeast corner located on the exact sampling point. d.

Clipping Procedures Different procedures are used to clip quadrats that are dry and those that are flooded.

Sampling Schedule During the sampling period do not collect all the samples in a cell at 1 time. Transects 25

Flooded Sites: the general procedure for harvesting flooded sites is the same as for dry sites except for the following: water depth (ern) is measured in tbe center of the quadrat; submersed vegetation and filamentous algae should be sampled within the water column by cutting along the boundaries of the quadrat and scooping the plants toward the middle when collecting

representative parts of the entire plant in proportion to tbeir abundance in the field. Ground samples are numbered consecutively and are given a prefix witb the type of sample and year, i.e. 82MAG89 for 82nd sample ground in 1989.

them. All fIlamentous algal mats should be collected separately and labelled "FIlAMENTOUS ALGAE". Field sampling time can be reduced to a minimum by using the following procedures for sorting samples. Place a couple of burlap bags around the site or in the boat prior to cutting the plants. One person should do all the cutting and should hand the clipped plants to a second person. The second person sorts the plants into appropriate piles on the burlap bags. When the sorting is completed the different fractions are placed into bags and labelled. Information from the field books should be entered on laboratory sheets (data sheet 23) at the end of each day.

Ground Sample Storage Samples should be stored in boxes labelled with tbe type of sample and year, ego MAG89.

Turn"Over Plots These plots are located in the 10 m buffer wnes witbin the experimental cells. The plots are to be monitored every 2 weeks beginning the first week of June. Only the dominant species (see aboveground macrophyte sampling) will be monitored.

Unknown Macrophytes If any dominant macrophyte is encountered that canDot be identified, it should be given a descriptive name, e.g. opposite leaved hairy herb, and a specimen collected and labelled the same way. This specimen should have flowers or fruits, if possible, and should be placed in a plastic bag along with the label. All unknown plants should be identified by MERP staff as soon as the samples are returned to the lab. Once the species are identified all bags of that species should be relabelled with the correct name. Each day a list should also be kept of all species that were collected as unknowns with their correct names. This list should be kept with the laboratory data sheets and voucher specimens of each unknown prepared.

First Count of Each Year

Count all green stems in each quadrat and mark each with orange flagging tape (to mark, loop tape very loosely around stem and staple in place). Record the number of marked stems under the "New Green Stems" column on the data sheet (data sbeet 24). Any stems tbat are dead standing stems from the previous year sbould be tagged with yellow flagging tape and listed under the DSL (dead standing litter) column. Record zeros in tbe remaining columns on tbe data sheet except for "Number of Collected Stems", and "Total Weight of Collected Stems" (see below). 2. Stem Classification During subsequent counts, stems must be classified as follows:

Drying Samples All samples should be placed in the drying room (80"C) and dried to a constant weight. If samples cannot be placed in the drying room immediately, they should be placed in the cold room (4"C). Weighing Samples Each sample is weighed to the nearest 0.01 gm. Sample weights are recorded on the lab sheet (data sheet 23). Grinding Samples All samples are ground in a Wiley mill. The general procedure is the same as with the belowground macrophyte samples. Before grinding, large samples are subsampled randomly, with care taken that entire plants are included in the subsample, not just plant parts. This ensures that the ground sample will contain 26

new green - unmarked stems

old green - green stems marked during previous counts (orange flagging tape)

new dead - stems tbat have died since the previous count (dead stems with orange flagging tape - see below)

old dead - stems tbat are still standing but were dead on previous counts (blue flagging tape)

vegetative - all non-flowering or non-fruiting stems witbin eacb of the above categories at tbe time of counting

flowering or frniting - all stems that are

flowering or fruiting within each of the above categories at the time of counting g. 3.

bag record: "TOP", species, date and stem type. Store these bags in a box labelled with TOP and the year (e.g. 1989 TOP).

dead standing litter from previous year (yellow flagging)

Rhizomes

for

Nutrient

Procedures Following the First Count During the collection of stems for biomass and nutrient analysis, collect about 500 gms. (fresh weight) of living rhizomes of each species. On return to the Station these rhizomes should be washed and then placed in the drying room (BO째C). Once dry, they should be ground and placed in Whirlpak bags labelled with: "TOP", date collected, species and "RHIZOMES". Store in the same box as the stem nutrient samples.

During all subsequent counts on the turn-over plots, count vegetative, flowering, and fruiting stems separately for each species (see data sheet 24). a.

Measure the water depth in the centre of all quadrats and enter the average on data sheet 24.

Count all standing stems marked with blue flagging tape and record as old dead stems. Remove the blue tape from any stems that have fallen down or been eaten off.

Collection of Determination

6. Belowground biomass should be determined for each species in June and September. Methods will follow MBG sampling.

Count all stems marked with orange flagging tape that are still green and record as old green

Photo Stations

stems.

Count all stems marked with orange flagging tape that are dead and record as new dead stems. Remove the orange tape from these stems and replace with blue tape.

Connt all unmarked green stems and record as new green stems. Mark these stems with orange flagging tape.

Count all dead standing stems from previous year (yellow tape). Remove tape from any stems that have toppled.

Permanent photo stations will be established on each cell (2 per cell) to provide a visual record of changes in macrophyte distribution and abundance. The observation tower overlooking each cell serves as 1 station. The second station is located in the centre of the south boundary of each cell. Cell 12 has only the tower photo-station. Colour slide photographs (35 mm) are to be taken at each station on the first day of each month from I May to I Nov. If weather conditions are not suitable, take the photographs as soon after these dates as possible. Three photos of each cell are to be taken from the observation towers. The first photo should include the north half of the cell, the second the centre of the cell, and the third the south half of the cell. Use the labelled brackets located on the tower windows. One photo is to be taken from the station on the south boundary of each cell. In the experimental cells, a pick-up truck is to be parked in the centre of the south dike of each cell. The photo is to be taken while standing in the back of the truck (to provide additional elevation). In cell 11, park on 22 Bay Road on the south end of the cell. There is no south photo-station on cell 12. Photos from these stations on the south boundary should include as much of the cell as possible. Careful records are to be kept during the photographing of the cells. Take 2 photos at all sites. This will result in 2 sets of pictures. After each picture is taken, record the picture number (on the camera) and the cell number. After developing, the cell number and date should be permanently marked on the frame of the slide.

Collection of Stems for Biomass and Nutrient Determination

At the time of each inventory, collect 50 living stems of each species in a randomly selected area adjacent to the quadrats. If the species is in flower or fruit, collect 50 vegetative stems and 50 flowering or fruiting stems. Place the collected stems in labelled burlap bags. Vegetative and flowering or fruiting stems should be placed in separate bags. Use the prefix "TOP" (turn-over plot) on the label, with species, date, and stem type (vegetative, flowering, and fruiting). On returning to the Station, place the bags in the drying room at BOoC until constant weight (2-3 days). Record the final dry weights of each stem type in the appropriate column on data sheet 24. Mter weighing, grind the entire sample and retain a subsample (approximately 20 gms) for later nutrient analysis. On the Whirlpak 27

1988b) or from year to year (van der Valk et aI. 1988).

General Vegetation Survey

Each August the vegetation is sampled in 30 1 x 1 m quadrats in each cell. Three quadrats are located at random along the work lanes in a cen. In each quadrat, the cover of each species and its standing titter are estimated to the nearest 5%. The mean height of each species and the water depth in the centre of the quadrat are measured as is the percentage of the shoots of a species that are flowering. From these data, the overall composition of the vegetation in the cells can be described and comparisons of the vegetation among cells made. For example, the frequency of occurrence of each species in a cell can be calculated as well as its total cover in a cell, and these data can be used to calculate the similarity of the vegetation in cells in different treatments in any 1 year as well as the similarity of the vegetation from 1 year to another

Literature Cited and Suggested Reading ANDERSON, M.G. 1978. Distribution and production of sago pondweed (Potomogeton pectinatus L.) On a northern prairie marsh. Ecology 59:154-160. GALINATO, M.l. AND AG. VAN DER VALK. 1986. Seed germination traits of annuals and emergents during drawdoWD in the Delta Marsh, Manitoba, Canada. Aquatic Botany 26:89-102. PEDERSON, R.L. 1981. Seed bank characteristics of the Delta Marsh: applications for wetland management. Pages 61-69 in Richardson, B., editor. Selected proceedings of the Midwest Conference On wetland values. Minnesota Water Planning Board - Branch, St. Paul, Minnesota. 66Opp.

in a given cell.

Permanent Quadrats PEDERSON, R.L. and AG. VAN DER VALK. 1984. Vegetation change and seed banks in marshes: ecological and management implementation. Transactions of the North American Wildlife and Natural Resources Conference 49:271-280.

Ten permanent quadrats were established in each cell early in the spring of the first year that they were drawndown (1983). These quadrats were located by dividing each cell in 5 equal wnes from north to soutb (lengthwise) and 2 equal wnes from east to west (widthwise). A permanent plot was placed at random in each of these 10 wnes. A permanent quadrat is 2 x 2 m. The comers and centre of a plot are marked by iron fence posts, and its perimeter and diagonals wired to delimit 4 triangular quadrats. The elevations of the plot at its 4 corners and centre were obtained with a level transit, and averaged to estimate the mean elevation of a plot. Permanent quadrats are sampled 3 times each growing season (June, July, and August). The number of flowering and vegetative shoots of each species is counted in the northern and southern triangular quadrats, i.e. 2 m' are sampled. The density of dead shoots of each dominant emergent species are also recorded. By using triangular quadrats, the vegetation in a permanent quadrat is never disturbed on 2 sides. During tbe drawdoWD years when seedling densities were very high, 1 m' subsamples were taken to estimate seedling density in a plot. Soil samples to determine soil moisture and salinity are taken adjacent to permanent quadrats. When they are flooded, water depth is also measured. Data from the permanent quadrats is used to monitor quantitative changes in the vegetation of an area over an elevation gradient (Welling et al. 1988a), during the growing season (Welling et aI.

Primary Production Processes, Part l. Pages 3-86 in Good, R.E., D.F. Whigham, and R. L. Simpson, editors. Freshwater wetlands: ecological processes and management potential. Academic

Press, New York, NY. 378pp. VAN DER VALK, A.G. 1986. The impact of litter and annual plants on recruitment of species from the seed bank of a lacustrine marsh. Aquatic Botany 24:13-26. VAN DER VALK, AG. 1988. From community ecology to vegetation management: providing a scientific basis for management. Transactions

of the North American Wildlife and Natural Resources Conference 53:463-470. VAN DER VALK, AG. (editor). 1989. Northern prairie wetlands. Iowa State University Press, Ames, Iowa. 4OOpp. VAN DER VALK, AG. and c.B. DAVIS. 1978a. Primary production of prairie glacial marshes. Pages 21-37 in Good, R.E., R.L. Simpson, and D.F. W!:igham, editors. Freshwater wetlands: ecological processes and management potential.

WALKER, J.M. 1%5. Vegetation changes with falling water levels in the Delta Marsh. Ph.D. Thesis. University of Manitoba, Winnipeg, Canada. 272pp.

Academic Press, New York, NY. 378 pp. VAN DER VALK, A.G. and C.B. DAVIS. 1978b. The role of seed banks in the vegetation dynamics of prairie glacial marshes. Ecology 59:322-335.

WELLING, C.H., R.L. PEDERSON, and A.G. VAN DER VALK. 1988a. Recruitment from the seed bank and the development of zonation of emergent vegetation during drawdown in a prairie marsh. Journal of Ecology 76:483-4%.

VAN DER VALK, A.G. and C.H. WELLING. 1988. The development of zonation in freshwater wetlands: an experimental approach. Pages 145-158 in During, HJ., M.JA. Werger and J .H. Williams, editors. Diversity and pattern in plant communities. The Hague, SPB Academic Publishing. 278pp.

WELLING, C.H., R.L. PEDERSON, and A.G. VAN DER VALK. 1988b. Temporal patterns in recruitment from the seed bank during drawdowns in a prairie wetland. Journal of Applied Ecology 25:999-1007.

WALKER, J .M. 1959. Vegetation studies on the Delta Marsh. M.S. Thesis. University of Manitoba, Winnipeg, Canada. 203pp.

Chapter 8

AERIAL PHOTOS AND COVER MAPS Patrick J. Caldwell' and Arnold G. van der Valk2 Introduction

Prints of each cell are made to a predetermined scale using a standardized template. By following this procedure, a uniform scale is maintained among cell, years and maps (contour and vegetation).

In order to determine changes in the location and area of the various vegetation types in the experimental cells, low-level aerial photographs are taken of each cell toward the end of each growing season. Vegetation maps are made from each aerial photograph and these maps are digitized by computer using a geographical information system. By manipulating the data, changes in the vegetation present in an area from year to year can be established and the areas of each vegetation type in a cell calculated. This information is used along with data from the primary production and decomposition studies to estimate total annual production in a cell and maximum amounts of carbon, nitrogen and phosphorus stored in the vegetation within the cell. Cover maps are also used to determine how much vegetation is being removed by muskrats and to evaluate the cells as habitat for waterfowl and other wildlife. In short, the aerial photographs and the vegetation maps made fro m them are essential for extrapolating the many different types of MERP data to entire cells.

Cover Mapping Immediately upon receipt of the aerial photograph prints, the cells should be cover mapped using mylar overlays on the prints. Mapping must be done in the same field season as the photographs are taken. AU conspicuous cover types larger than 100 m' should be mapped. Each cover should be traced and labelled on the mylar and identified by ground truthing. Dominant emergent and understory species are to be recorded on the mylar overlay. During ground-truthing, 5 water depths are to be measured within each continuous dominant cover type. The measurements are to be taken at locations within the cover type to represent as nearly as possible both shallow and deep portions of the area. Each measurement site should be located as precisely as possible on the cover map overlay by utilizing visible land or vegetation features clearly discernible on photos and on the ground. The water level at the staff gauges within each cell is also to be recorded on the same day. Water depth measurements will then be converted to ground elevations by referencing them back to the staff gauge readings. Final cover maps are to be traced in ink from the overlay onto 216 x 279 mm mylar. The maps are then digitized on computer using a dot grid method developed by the Soil Science Department at Iowa State University. Contour maps using a 10 em interval were also produced early in the study. These have also been digitized and can be overlaid on the vegetation maps to allow determination of vegetation changes according to contour levels and water depths throughout the study.

Aerial Photographs Aerial infrared photographs are taken annually of each cell within the experimental complex. Photography is completed between mid-morning and noon on a day with no cloud cover between 6 and 20 August. All cells are photographed on the same day. Each photo is taken with 1 cell centered within the frame. The cells are photographed with a 70 mm camera from a height of 610 m above ground level or 854 m above sea level. The camera is equipped with a 50 mm lens corrected for aerial photography with a Wrattan B and W 16 orange filter.

, Ducks Unlimited Canada, 1190 Waverley Street, Winnpeg, Manitoba R3T 2E2 , Department of Botany, Bessey Hall, Iowa State University, Ames, Iowa 50011 30

Chapter 9

DECOMPOSITION Henry R. Murkin', Arnold G. van der Val~ and Jeffrey W. Nelson'

Introduction

dried to constant weight (ca. 48 h) at ZOOC. This was recorded as fresh weight. To obtain "time zero' nutrient analyses, random control samples for each species were removed, dried to constant weight at BOoC, ground in a 4O-mesh Wiley mill, and stored in labelled Whirlpak bags. To estimate a weight ratio between fresh litter (ZOOC constant weight) and dry litter (BOOC constant weight), 30 random preweighed fresh litter (2O"C constant weight) subsaDlples per species were dried to 80"C constant weight. Weights are recorded to 0.01 g accuracy, data sheet 25. Litter bags were prepared by placing predetermined amounts of fresh litter (see below) in sealed 20 x 40 em polyethylene mesh bags (1 mm mesh size) . One-hundred fifty litter bags were prepared for each species. Each bag was numbered with a plastic tag. Tag numbers are:

The high pnmary productivity in wetlands ensures that plant litter production and decomposition are important factors in wetland nutrient budgets. There are 3 components of the litter pool in prairie wetlands: standing litter, fallen litter, and dissolved organic compounds that leach from both standing and fallen litter (van der Valk and Davis 1978a). Material enters the standing litter compartment with the death of the leaf or shoots. In prairie marshes, litter is transferred from the standing litter compartment to fallen litter primarily through fragmentation by wind, snow, and ice action. In areas of high muskrat activity, feeding and house building can result in living plant tissues directly entering the fallen litter stage. Once within the fallen litter compartment, decomposition rates increase due to the growth of bacteria, fungi, and other consumers on the litter particles. The MERP decomposition studies are a series of short-term studies eXaDlining the decomposition rates of both above- and belowground components of the dominant emergent species and the transfer of litter from the standing to the fallen litter compartment within the experimental cells.

751 - 900 Phragmites australis 901 - 1050 Scolochloa [estueaeea 1051 - 1200 Typha glauca 1201 - 1350 Scirpus maritimus 1351 - 1500 Scirpus lacustris Initial shoot weights for the 5 species are approximately 15 g for P. australis, and T. glauea, and 10 g for S. [estueaeea, S. maritimus, and S. laeustris. Each bag was weighed to the nearest 0.01 g before and after filling.

Shoot Decomposition Study (1986) The purpose of this study is to determine the decomposition rate of the 5 dominant emergent species from the 1985 growing season: Phragmites australis, Scirpus iacustris, Scirpus maritimus, Scolochloa [estueaeea, and Typha glauea. 1.

Litter Bag Deployment

Five bags of each species were deployed at 3 different sites in each cell according to the schedule in Table 1, beginning on 26 May 1986. Five bags of a single species were attached to individual wire tethers (2 m long) and secured to a 5 x 5 cm marker post at each site. Bags were allowed to sink if nooded, otherwise they were placed nat on the marsh substrate.

Litter Bag Preparation

Shoots of the 5 species were collected on 23 April 1986. Only portions above the water surface were collected. Each species saDlple was a composite from a number of cells. SaDlples were

Delta Waterfowl and Wetlands Research Station, R.R. 1, Portage la Prairie, Manitoba R1N 3Al

Department of Botany, Bessey Hall, Iowa State University, Ames, Iowa 50010

Ducks Unlimited Inc., One Waterfowl Way, Long Grove, Illinois 60047

Litter Bag Collection Each bag was weighed to the nearest 0.01 g before and after filling (data sheet 26).

The collection schedule is 1 litter bag per site (150 total) during each of the 5 collection periods: 48 h after deployment, July 1986, July 1987 and June 1988. See Table 1 for sampling order. To remove bags, follow the wire tether to the substrate surface and carefully lift the bag and cut the wire. In the lab, the litter is removed from the bag and washed in distilled water to remove soil and extraneous plant and animal material. Record tag number and keep tag with the sample. Samples are dried to constant weight (ca. 48 h) (8O"C) and weighed. Weights (to 0.01 g) are recorded. Following weighing, each litter sample is ground in a 4O-mesh Wiley mill and stored in a labelled Whirlpak bag for nutrient analyses (N,P,C). The % dry weight, N, P, and C remaining will be calculated for each bag based on values from the initial control samples.

Five bags of each species were deployed at 3 different sites in each cell according to the schedule in Table 1, beginning in May 1988. Five bags of a single species were attached to individual wire tethers (2 m long) and secured to a 5 x 5 em marker post. Bags were inserted below the soil surface in slits created by inserting a spade and pulling it to the side. Bags were positioned so that the shoot protruded above the marsh substrate. The criteria for site selection was to represent the natural depth range for each species as observed during the 1987 growing season. 3.

The purpose of this study is to determine the decomposition rate of roots and rhizomes from 4 dominant emergent species in the Delta Marsh: Scolochloa lestueaeea, Scirpus laeustris, Phragmites australis, and Typha glauco. Litter Bag Preparation

Roots and rhizomes of the 4 designated species were collected in April of 1988. A 5-cm segment of shoot was left attached to each sample. Each species sample was a composite from a number of cells. Samples were dried to constant weight (ca. 48 h) at 20째e. This was recorded as fresh weight. To obtain "time zero" nutrient analyses, random control samples of each species were removed, dried to constant weight at 8O"C, ground in a 4O-mesh Wiley mill, and stored in labelled Whirlpak bags. The ratio between fresh litter (2O"C) and dry litter (80째C) was determined by taking 30 random prcweighed fresh litter subsamples per species and drying them to 8O"C constant weight. Weights are recorded (to 0.01 g) on data sheet 25. Litter bags were prepared by placing 10 g fresh weight in sealed 20 x 40 cm polyethylene mesh bags (1 mm mesh size). One-hundred fifty litter bags were prepared for each species. Each bag received a plastic tag. Tag numbers are:

Litter Fall Study (time series experiment) 1.

Matching Stems 1986-1989

The purpose of this study is to estimate changes in aerial litter standing crop (biomass and (N,P,C)) by monitoring seasonal litter fall patterns of Typha glauca, Scirpus lacustris, Seholochloalestucacea, and Phragmites australis. a.

1501 1651 1801 1951

Litter Bag Collection

The collection schedule is 1 litter bag per site (150 total) during each of the following collection periods: 48 h after deployment, July 1988, September 1988, June 1989 and August 1989. The removal of bags is accom plished by following the wire tether to the substrate surface, and after cutting the wire, carefully removing the bag from the substrate. In the lab, the litter is removed from the bag and washed in distilled water to remove soil and extraneous plant and animal material. Record tag number on data sheet 26 and keep tag with the sample. Samples are dried to constant weight (ca. 48 h) at BO'C. Weights (to 0.01 g) are recorded on data sheet 26. Following weighing, each sample is ground in a 4O-mesh Wiley mill and stored in a labelled Whirlpak bag for nutrient analyses (N,P,C). As with the shoot study, % dry weight, N, P, and C remaining will be calculated for each bag based on values from the initial control samples.

Belowground Decomposition Study (1988)

Litter Bag Deployment

- 1650 S. lestucacea - 1800 S. lacustris - 1950 P. australis - 2100 T. glauca

Field Procedures Aerial standing live shoots are marked for this experiment. Changes in the aerial standing litter component are then measured over time. Replications are part of a time series based on

the initiation date: Experiment 1 - May 1986, Experiment 2 - August 1986, Experiment 3 August 1987, Experiment 4 - August 1988. To initiate each experiment, 4 live shoots are chosen at each of 70 sites. The criteria for site selection is to represent the natural depth range for each species. New sites are selected in consecutive years when vegetation becomes sparse or void on previously selected sites. The only species to be marked at each site is that of the assigned cover type. Each of the 4 shoots is marked with a different colour of floating flagging tape. Water depth is recorded. These shoots are followed until they have completely toppled. They are used as a reference when collecting matching stems for weight and nutrient analyses. The flrst collection of matching stems is concurrent with the initial marking of permanent stems; i.e. August for aU but the May 1986 experiment. Further collections of the May 1986 experiment were Further June, July and October 1986. collections of the August 1986, 1987, and 1988 experiments are October of that year (prior to ice in), and the following May, June, July, and September. A nearby stem, matching the marked stems as closely as possible in length, thickness, and number of leaves, is clipped i) at the substrate surface (dry sites) or ii) at the water surface (also clip off leaf portions that are submersed) (flooded sites). Record water level in the centre of the 4 marked stems on each sampling visit. Generally, each experiment has a time frame of 1 year from date of initiation. Aerial standing litter (dead) is assumed to be from the previous growing season. An exception is Phragmites which persists for 1 + years and should be separated as fresh litter (shiny; tan colour) or old litter (dull; grey colour) when stems are matched. b.

litter biomass and nutrient concentration (N,P,C) using clip quadrats.

Lab Procedures Place matched stems in separate paper bags and return to the lab for drying and weighing. Bags should be labelled with species, date of collection, site #, flagging colour, cell #, and "Matching Stems路 and the year and month the study was initiated. All litter samples should be immediately dried to constant weight at 8O"c. Record data on data sheet 27.

2. Clip Quadrats The purpose of this study is to estimate annual standing litter production by monitoring standing 33

Field Procedures A total of 150 sites (15 sites in each of the 10 diked MERP cells) are selected. The criteria for site selection is to represent the natural depth range for each species. New sites are selected when vegetation becomes sparse or void on previously selected sites. Each site is marked by a numbered stake. Sampling requires that 1 x 1 m quadrats be clipped at aU sites during May and August. A randomly selected, undisturbed plot is located within 2 m of the site marker stake. All data is recorded on data sheet 28. Month of sampling and quadrat moisture condition (dry vs. flooded) determine the sampling procedure at each site (Fig. 9.1). Water depth is recorded at the centre of each quadrat. If a plant is rooted within the quadrat all foliage is included with the sample even if extending beyond the quadrat bounds. Within a quadrat aU dominant emergent species (i.e. Typha glauco, Phrogmites australis, Carex atherodes, Scirpus lacustris, Scirpus maritimlls) are treated separately. Other species with < 25% cover are combined and labelled "MINOR SPECIES". After clipping, plants are sorted and placed into separate paper bags and labelled with 1) species name, 2) collection date, 3) site number, 4) cell,S) moisture condition and 6) litter type (i.e. dry standing litter, aerial standing litter, aerial standing live shoots, flooded standing live shoots). Aerial standing litter is assumed to be from the previous year with 1 exception Phragmites australis. Phragmites can persist for 1 + years and should be separated as fresh litter (shiny; tan color) or old litter (dull; grey color).

Lab procedures All litter samples should be immediately dried to constant weight (8O"C) and weighed to 0.01 g accuracy. All dried samples are ground in a 4O-mesh Wiley mill and each stored in labelled Whirlpak bags. A single sample which occupies more than 1 storage bag will need to be subsampled (20 g subsample). The Whirlpak bags are labelled with a 2 part code consisting of an appropriate preflx and a unique number, i.e. Aerial89 9506. The code is recorded on data sheet 28 (clip quadrats).

NECKLES,

and in a Whitetop marsh. Unpublished M.S. Thesis, Univ. of Minnesota, Minneapolis, MN. l09pp.

Literature Cited and Suggested Readings

H.A.

1984.

Plant

macroinvertebrate responses to water regime

DAVIS, c.B.AND AG. VANDER VALK. 1978a. The decomposition of standing and fallen litter of Typha glauca and Scirpus fluviatilis. Can. J. Bot. 56:662-675.

POLUNIN, N.V.C. 1984. The decomposition of emergent macrophytes in fresh water. Adv. Ecol. Res. 14:115-166.

DAVIS, C.B. ANDAG. VANDER VALK 1978b. Litter decomposition in prairie glacial marshes. Pages 99-113 in Good, R.E., D.F. Whigham, and R.C. Simpson, editors. Freshwater wetlands: ecological processes and management potential. Acad. Press, New York, NY. 378pp.

WEBSTER, J.R. AND E.F. BENFIELD. 1986. Vascular plant breakdown in freshwater ecosystems. Ann. Rev. Ecol. Syst. 17:567-594. WIEDER, RK, AND G.E. LANG. 1982. A critique of the analytical methods used in examining decomposition data obtained from litter bags. Ecology 63:1636-1642.

GODSHALK, G.L. AND R.G. WETZEL. 1978. Decomposition of aquatic angiosperms. II. particulate components. Aquat. Bot. 5:301-327. MURKIN, H.R. 1989. The basis for food chains in prairie wetlands. Pages 316-338 ill van der Valk, A.G., editor. Northern prairie wetlands. Iowa State Univ. Press, Ames, IA. 4OOpp.

Fig. 9.1 Clip Quadrat Sampling Flow Chart Month of Collection Table 1. Schedule based on a 5 day cycle for deployment and collection of litter bags and standing litter clip plots

August

May Day

Within Cell DayÂˇ Number

Moisture ---Dry--- > Condition 1

1 2

2 5

high medium

1 2

7 8

low - 1 yr drawdown low - 2 yr drawdown

1 2

1 6

medium high

1 2

3 4

low - 1 yr drawdown low - 2 yr drawdown

1 2

9 10

medium high

Treatment

Clip

< --- Dry--- Moisture Condition

1. All Standing Litter

Flooded

I I

Clip

1. Aerial Standing

Litter

â&#x20AC;˘ Assumes 30 sites (15/cell) can be visited within a morning and samples collected can be processed that afternoon. Sam pies can be weighed the following week.

1. Aerial Standing

Litter 2. Aerial Standing Live Shoots 3. Flooded Standing Live Shoots

Chapter 10

INVERTEBRATES Lisette C.M. Ross and Henry R. Murkin' Introduction

invertebrate habitats to be sam pled: the water column and substrate. Because no 1 sampler effectively samples the entire invertebrate community (Swanson and Meyer 1973), a variety of samplers are used to provide separate data on the' nektonic, benthic, and epiphytic groups.

Aquatic invertebrates are an important component of freshwater wetland systems, yet we know little about their life histories and general ecology. They serve as critical food resources for a wide range of marsh wildlife (Murkin and Batt 1987), however, their role in wetland nutrient dynamics and macrophyte decomposition require further investigation. Determination of seasonal differences in aquatic macroinvertebrate populations within and between the dominant vegetation types and the various water level treatments within MERP will be a significant step in improving our understanding of their ecology and function in wetland systems.

Activity Traps

Activity traps (Fig. 10.1) are used at sampling sites with standing water to provide an index to the population levels of free-swimming invertebrates (Murkin et aI. 1983). Activity traps are set biweekly for a 24-h period. 2.

Sampling Design and Samplers

Artificial Substrates

An artificial substrate (Fig. 10.2) is placed at each sampling station with standing water to provide an index to Gastropodinae and Chironomidae populations colonizing submersed surfaces. They are to be sampled on the same schedule as activity traps.

Invertebrate sampling stations within each experimental and control cell were stratified by cover (vegetation) type based on the dominant emergent plants: cattail (Typha glauca), hardstem bulrush (Scirpus lacustris ssp. glaucus) softstem bulrush (Scirpus lacustris ssp. va/idus), whitetop (Scholochloa festucaeea), rayless aster (Asler brachyaclis), Phragmites (Phragmites australis), and open water (standing water with no emergent vegetation). Two water depths (30 cm (shallow) and 60 ern (deep禄 strata were also determined for every cover type. For each cover type 6 sampling stations were randomly chosen for each water depth. Sampling site locations will remain the same each year to allow comparisons over time. The number of samples per station were determined from calculations of Downing (1979) using information on expected invertebrate densities (Murkin et aI. 1982) and sampler size. Density, biomass, and number of taxa will be used as response variables. Due to taxonomic problems with the invertebrates of wetlands and the overlap of the trophic categories suggested by Cummins (1973), 2 broad trophic levels will be distinguished; herbivore-detritivore and predator路 parasite. At each sam piing station, there are 2

Emergence Traps

Emergence traps (Fig. 10.3) are used to provide an index to aquatic insect population levels within the cells. Most invertebrates within the benthos of the MERP cells are larval stages of aquatic insects. However, there are also aquatic insects associated with submerged vegetation, which cannot be sampled by a simple substrate or core sampler. The emergence trap used in this study is a modification of LeSage and Harrison's (1979) model "Week" (see Wrubleski (1984) for trap modifications). The base size of the trap is 0.5 m . Traps will be monitored on a weekly basis. Field And Lab Procedures

Activity Traps

Setting Activity Traps Slowly submerse the activity trap jar. Attach the funnel over the mouth and suspend the trap

, Delta Waterfowl and Wetlands Research Station, R.R. 1, Portage la Prairie, Manitoba R1N 3A1 35

from the stake so that it lies in a horizontal position midway in the water column. There should be no air bubbles present in the activity trap once set, as this will allow the survival of air - breathing predaceous insects in the trap. Maximum-minimum thermometers are set the same time as activity traps. The thermometers used in this study have metal floats on top of the mercury column that are reset with the magnet provided. Thermometers should be reset under water and fastened midway in the water column. b.

Collecting Activity Traps To collect the activity trap sample, carefully raise the trap to the surface. At the surface immediately tilt the trap upright. Remove the funnel and pour the trap contents through a standard U.S. #35 sieve. Wash the contents of the sieve into a sample jar labelled with site, date, maximum and minimum temperatures. When reading water temperatures, remove the thermometer from the stake and raise it to just below the water surface. Read the temperatures while the thermometer is completely submersed. Record the water depth. If activity trap samples are not sorted immediately after returning from the field, refrigerate samples at 4째C until sorted.

Sorting Activity Traps Identify all macroinvertebrates in the sample to the taxonomic level necessary to determine the trophic level of the organism, and record total numbers of each taxon. In most cases, identification to family is sufficient. Notable exceptions occur in the Cbironomidae, which must be identified as Tanypodinae (Predators) or Non-Tanypodinae (all other subfamilies combined; Herbivore-detritivore). For many of the lower invertebrate groups, higher levels of classification suffice (e.g. Oligochaeta, Nematoda). Check the MERP reference collection for aid in identification and classification. Many invertebrates must be further classified for size (small, medium, large) and life stage (larvae, pupae, adult). Cheek the list available in the MERP lab to see which families are currently divided into size classes and life stages and be sure to include this information in your identification. Any invertebrates that cannot be identified, or are different from the size classes in the reference collection should be placed in a separate vial labelled with date, sample type, site, and any comments (Unknown

A, B, C, etc.; smaller than size classes provided, etc.). Following counting and recording of information on data sheet 29, select 7 to 10 macroinvertebrate representatives from the sample and place in a labelled vial fIlled with 70% alcohol. These representatives will be kept for future reference.

Use of Subsampler To facilitate counting large numbers of individuals of a single taxon (e.g. Cladocerans), the mechanical subsampler should be used. The subsampler used in this study is designed after Waters (1969). After minor taxon in the sample have been counted, turn the subsampler on and rinse the sample through the funnel on top. Your sample will be distributed equally into 8 collecting sieves in the subsampler. Count the individuals in only as many sieves as necessary to total at least 50 individuals. Divide into 8 the number of sieves looked at and multiply by the number of individuals counted to obtain the total number of individuals in the sample (e.g. if it was necessary to count Cladoceran in 2 sieves to have counted at least 50 Cladocerans, and 60 were counted in those 2 sieves, then (8/2x(6O)=240 c1adocerans in sample).

Artificial Substrates

Setting Artificial Substrates One artificial substrate is placed at specific sampling stations at the beginning of the field season. They are sam pled at the same time as activity traps.

b. Collecting Artificial Substrates Carefully scrape all visible surfaces with a flat scraper and place contents into a collecting jar labelled with the site and date. Suspend it midway in the water column. Do not push it into the bottom substrate. If sam pies are not sorted immediately after returning from the field, refrigerate the samples at 4째C until sorted.

Sorting Artificial Substrates Identify and size class all Gastropodinae to family level and all Chironomidae to Tanypodinae or non-Tanypodinae. All those identified should be kept for future reference. Record data on data sheet 30.

Emergence Traps

Collecting Emergence Traps

mean weight of > 1.0 mg these groups no longer have to be saved for dry weight determination.

To sample an emergence trap, remove the sample jar from the trap and loosely screw a new jar onto the head. Place a lid on the sample jar removed and label with sample site, date, water depth, water temperature, substrate surface temperature, and substrate temperature. When setting and removing samples from emergence traps the fonowing should be checked and fonowed:

Literature Cited and Suggested Readings CAIRNS J., Jr., editor. 1982. Artificial substrates. Ann Arbor Science Publishers Inc., Ann Arbor, MI. 279pp. CUMMINS, K.W. 1973. Trophic relations of aquatic insects. Ann. Rev. Ent. 18:183-206.

I. Tie-downs: must be secure in flooded sites. Ii. Plastic heads must be clean inside (remove

DOWNING, JA. 1979. Aggregation, transformation and the design of benthos sampling programs. J. Fish. Res. Board Can. 36:1454-1463.

and save invertebrates stuck to spider webs) and out (clean off any bird droppings or other accumulated dirt). Cut any vegetation growing into the plexiglass head. Check for broken heads and replace if necessary. ill. Net and wire skirt: if damaged replace trap. iv. Trap bottom: be sure base is undamaged and clear of algae. Bottom tubing should be flush with the substrate or water surface. b.

LESAGE, L. and A.D. HARRISON. 1979. 1m proved traps and techniques for the study of emerging aquatic insects. Ent. News 90:65-78. MURKIN, H.R. 1983. Responses by aquatic invertebrates to prolonged flooding of marsh habitat. Ph.D. Thesis, Utah State Univ., Logan, UT.112pp.

Sorting Emergence Traps All Chironomidae wiD be identified to subfamily level. Of the remaining Diptera, 12 families are recognized; Ceratopogonidae, Chaoboridae, Culicidae, Dolichopodidae, Ephydridae, Psychodidae, Scathophagidae, Sciomyzidae, Stratiomyidae, Syrphidae, Tipulidae, and Tabanidae. All other Diptera should be combined as "Other Diptera". All other orders are taken to family except Hymenoptera, Lepidoptera, Hemiptera, and Homoptera for which order is sufficient. Helodidae is the only family of Coleopterans identified. All other Coleoptera wiD be counted and noted as "Other Coleoptera". Record information on data sheet 31.

MURKIN, H.R. and B.D.J. BATT. 1987. The interactions of vertebrates and invertebrates in peatlands and marshes. Pages 15-30 in Rosenberg, D.M. and H.V. Danks, editors. Aquatic insects of peatlands and marshes in Canada. Memoirs of the Entomological Society of Canada 140. MURKIN, H.R., and JA. KADLEC. 1986. Responses by benthic macroinvertebrates to prolonged flooding of marsh habitat. Can. J. Zool. 64:65-72.

Dry-Weight Determinations

MURKIN, H.R., R.M. KAMINSKI, and R.D. TITMAN. 1982. Responses by dabbling ducks and aquatic invertebrates 10 an experimentally manipulated cattail marsh. Can. J . Zool. 60:2324-2332.

To allow determination of biomass, a representative sample of all invertebrate taxa and size classes should be kept for dry-weight determination. Use aluminum foil pans which have been previously dried overnight (10S'C for 24 h) and weighed. Keep a tally of the number of individuals in each pan (see data sheet 32) before drying the pans at 105'C overnight. These should be weighed the following day, and Ihe weights of the pans and invertebrates recorded. Use tongs for handling the pans, i.e. removing pans from the oven (empty or full), cooling them in the desiccator, and weighing them. After collecting 500 individuals of any taxon or size class which has a mean weight of < 1.0 mg and 200 individuals in any taxon having a

MURKIN, H.R ., P.G. ABBOTT, and JA. KADLEC. 1983. A comparison of activity traps and sweep nets for sampling nektonic invertebrates in wetlands. Freshw. Invert. BioI. 2:99-106. NECKLES, HA. 1984. Plant and macroinvertebrate responses to water regime in a whitetop marsh. M.S. Thesis, Univ. of Minnesota, St. Paul, MN. l09pp.

RESH, V.H. and D.M. ROSENBERG. 1984. The ecology of aquatic insects. Praeger Publishers, New York, NY. 625pp.

WATERS, T.F. 1969. The turnover rate in production ecology of freshwater invertebrates. Am. Nat. 103:173-185.

SWANSON, GA. and M.I. MEYER. 1973. The role of invertebrates in the feeding ecology of Anatinae during the breeding season. Pages 143-185 in Waterfowl Habitat Management Symposium, Moncton, NB. 306pp.

WRUBLESKI, DA. 1984. Species composition, emergence phenologies, and relative abundances of chironomidae (Diptera) from the Delta Marsh, Manitoba Canada. M.S. Thesis, Univ. of Manitoba, Winnipeg, MB. 115pp.

Fig. 10.1 Activity Trap

Fig. 10.2 Artificial Substrate

Fig. 10.3 Emergence Trap

Chapter 11

VERTEBRATES William R. Clark' and Henry R. Murkin' Muskrats 1.

population density require mark-recapture experiments (Clay and Clark 1985, Kroeker 1988), especially if there is a desire to measure other parameters such as survival and dispersal.

Background and Objectives

Muskrats can recolonize marshes that have been reflooded following drawdoWD within 1 year and reach substantial densities within a few years (Errington 1%3, Perry 1982). Kroeker (1988) estimated that populations in MERP cells increased from an average prebreeding density of 2/ha in May 1986 to an average fall density of 21/ha in October 1987. Muskrat populations exhibit inverse density dependence in both reproductive and survival rates (Errington 1%3, Clay and Clark 1985, Clark 1987, Kroeker 1988), although it has not been clearly shown to what extent changes in these parameters are responsible for overall population trends during recolonization. In addition it is difficult to separate dispersal and mortality within estimated disappearance rates. Water depth is considered critical in determining vegetation characteristics of wetlands and habitat use by muskrats (Lay and O'Neil 1942, Belliose and Low 1943, Errington 1%3, Palmisano 1972, Perry 1982, Proulx and Buckland 1986), yet nO quantitative data are found in the literature which relate minimum water depth and habitat condition to overwinter

Sampling Rationale and Analysis

Muskrat density will be estimated by sampling for 6 days using capture-recapture techniques. Estimates of population size will be calculated assuming a closed population (Otis et al. 1978). The goal is to determine 2 point estimates and associated variances for each cell each year, 1 in May and another in September. Survival rates will be estimated from the capture-recapture data using a hybrid design described by Pollock (1982). Captures from the 6day periods will be pooled and Jolly-Seber (Seber 1982:199-202) estimates of survival rate between the point estimates of population size will be calculated using POPAN-2 (Arnason and Baniuk 1978). Densities, survival rates, rates of increase, and changes in body condition will be compared among treatments and seasons using analysis of variance with an error term of variation among replicates within a treatment. Associations between density and the other variables will be tested with simple, linear regression. Chi-square tests will be used to identify differences in movements among cells.

survival rates. The objectives of monitoring muskrat responses within each cell are i) to document population recolonization and to estimate density within each water level of the MERP experiment and ii) to determine which demographic parameters are responsible for the observed density changes. The expectation is that muskrat populations will be most dense and have the greatest survival during winter in cells with deepest water. Muskrat populations have been monitored by house counts (Dozier 1948, Mathiak 1966), vegetation indicators (McCabe 1983), and harvest statistics (Perry 1982) but such approaches provide only an imprecise index of population trend (Clark and Andrews 1982). Accurate estimates of

Population Estimation

Trapping periods will begin shortly after ice-out on all cells and all cells should be completed in about 6 weeks. Trapping resumes approximately 1 September to allow ample time for completion before freeze in the fall. Each cell is trapped for 6 consecutive days and traps are moved to a new cell on the seventh day. If severe cold or windy weather occurs, it is preferable to delay the start of a new experiment although it is acceptable to close the traps for a day or 2 as long as 6 full days are completed. Cells should be trapped in random

'Department of Animal Ecology, 124 Sciences Hall II, Iowa State University, Ames, Iowa 50011 'Delta Waterfowl and Wetlands Research Station, R.R. 1, Portage la Prairie, Manitoba RlN 3A1 39

order, usually trapping 2 cells simultaneously. Each cell is set with 45 15 x 15 cm, single or double·door, Tomahawk live traps, arranged in a grid along the 10 work lanes. Because cell 2 is somewhat wider than the other cells, 55 traps are set to keep trapping effort constant among all cells. Five trapping stations are designated in each work lane; 4 are located within the celi whereas the fifth is located on the west side ofthe borrow ditch. Cell 2 is the exception and has 5 traps located within the cell instead of 4. The traps are numbered beginning in the northeast corner of the cell. The first trap in lane 1 is number 1 and the last trap in lane 1 (the trap on the west side of the borrow ditch) is number 5. The first trap in lane 2 (beginning at the east end) is number 6 and the last trap in lane 2 (across the borrow ditch) is number 10, etc. Because of the larger size of cell 2, the numbering system has a slight variation. Instead of trap numbers 5, 10, 15, etc. being located along the borrow ditch, they are in the ceU. Therefore the traps along the borrow ditch are numbered 51 to 60 from the north end to the south end of the ditch. The 4 traps (5 in cell 2) along the worklane in the cell should be spaced evenly. The first trap should be 15·20 m (about 20 steps) from the base of the east dike, and subsequent traps are spaced 35·45 m apart (about 40·50 steps). This spacing should result in the fourth trap about 15·20 m cast of the borrow ditch. The goal is to get systematic coverage of the entire cell (dry areas of the cells included). The traps west of the borrow ditch are set on the area between the base of the dike and the borrow ditch. They should be set over water whenever possible to minimize the possibility of predation by raccoons and mink. Because the borrow ditch is an easy travel route for muskrats, the borrow ditch traps are likely visited more frequently by muskrats than the other traps. In an attempt to compensate for this chance and to keep trapping effort constant, only every other lane along the borrow ditch is set with a trap. New trap locations in the cells are established each fall to accommodate the shifting work lanes. At the time the new trap sites are located, it is determined by the toss of a coin whether the odd or even trap sites along the borrow ditch will be used. The same locations are used again the next spring. When choosing new trap sites, take into account the relative location of adjacent work lanes. For example, if work lanes 2 and 3 are only a few meters apart, set each of the sites in lane 2 a few meters north of the lane and the sites in lane 3 a few meters south of the lane to accomplish even coverage of the cell. Traps are suspended above the water from 2

or more pieces of lath. The traps are tied to the lath with plastic bailing twine. The twine should be tied to opposite upper corners of the trap so that captured muskrats cannot reach them with their teeth. The floor of the trap at the door should be at, or slightly below, the water line (only the main door is used on double door traps). The other end of the trap should be above the water to allow the captured animals to remain dry. No bait is used, but it is extremely important to cover the traps with ample vegetation so they appear as small muskrat shelters. The vegetation covering the trap is the only protection the muskrats have from the cold, wind and rain as well as from the hot sun. Traps on dry land need not be tied to 2 pieces of lath but should be adequately covered with vegetation. The trap site number should be marked on at least 1 of the pieces of lath and the location should be well flagged with marking tape. After the trap has been set and covered, test the trigger mechanism to ensure it is not blocked by vegetation, twine or lath and that it is easily tripped. Traps should be checked as early as possible each morning to minimize stress to captured individuals. 4.

Handling Captured Animals

It is easiest to carry an extra trap of known weight to use as a handling cage. Set the handling trap and carefully transfer the captured animal from the trap to the handling trap. The animal can be held in the handling trap while the trapsite is reset and recovered. Weigh the animal to the nearest 25 g while it is still in the handling trap (remembering to subtract the weight of the trap). Periodically check the tare weight of the scale to ensure its accuracy. An animal is classified as a breeding adult if the weight exceeds 750 g (Kroeker 1988), otherwise it is a juvenile of the year. In spring, all animals are classified as breeding adults regardless of weight. A handling cone (McCabe 1983) is used to remove animals from the trap and to restrain them. Be sure to check the cones for weak strings or bent wires that will allow the animal to escape through the sides. Muskrats can squeeze through spaces 3 cm wide. Place the sleeve of the handling cone over the end and carefully transfer the animal to the cone from the handling trap. Slightly tipping the trap, extending the cone, or gently blowing at the animal are useful in directing it into the cone. If the animal should escape into the canoe, set the handling trap, place it along 1 side of the canoe bottom, and use the paddle to direct the animal into the trap. The paddle is also useful to keep the escaped muskrat from jumping out of the canoe. Remember, an

escaped, unmarked, unidentified animal provides no information. Once in the cone, animals will usually relax but be beware of the exception! A paddle placed across the thwarts of a canoe makes a convenient work platform. Animals should be marked immediately with identically-numbered #1 monel fIsh tags in each ear. Be sure to place the tag at the base of the rear edge of the ear so that it pierces the cartilage and the number is visible from the back of the ear. This facilitates reading the tag upon recapture. If 1 tag of a pair is accidently destroyed, discard the extra and use the next pair of tags. If 1 tag has already been applied, use a tag from the next pair of tags, discard the extra, and record the tag numbers as they occur. Always make sure the animal has 2 ear tags to protect against possible loss of marks. Sex the animal by external characteristics (Larson and Taber 1980:163). The urinary papilla of the female and the sheathed penis of the male are externally similar in appearance. However, it is possible to expose the penis of the male by stripping back the prepuce, ensuring positive identifIcation. Additionally, the female has a bare, sometimes closed, vaginal opening at the posterior base of the urinary papilla. The paired testes of the male, although located internally, are very noticeable in the spring. By fall, they have atrophied and are no longer useful as an identifying characteristic. Measure the body length from nose to the base of the tail (where the bare, scaly skin starts) to the nearest 5 mm. Try to measure the animal when it is relaxed and stretched out in the cone. Measure the hind foot from heel to longest toe to the nearest 1 mm .

days on which the animal was not captured, ie. left justify all recapture data. For example, an animal whose history was captured on day 1 in trap 10, recaptured on day 3 in trap 11, and recaptured on day 4 in trap 11 should be recorded as: 01 10,03 11,04 11. If an animal is dead in the trap, record the occasion as a negative number, ie. dead on day 2 is recorded as -2. Birds 1.

A wide variety of birds use wetland habitats for all or part of their life cycles (Weller 1981). The abundant food supplies and the normally heterogeneous nature of the habitats play roles in the high avian use and productivity in these environments. Invertebrate food resources have been shown to be an important factor in determining waterfowl and blackbird use of prairie wetlands, particularly during the breeding season (Murkin 1979, Murkin et al. 1982, Murkin and Kadlec 1986, Murkin and Batt 1987). Habitat structure, especially cover-water ratios, have also been shown to influence avian use of these habitats (Weller and Spatcher 1%3, Weller and Fredrickson 1974, Kaminski and Prince 1981, Murkin et al. 1982). The MERP experiment will provide an unprecedented opportunity to examine the effects of a wide range of enviromnental factors on the avian use of the cells during the experimental wet-dry cycle. 2. Weekly Census

Regular weekly counts will be made from 1 May to 30 June for blackbirds and from 1 May to 30 October for waterfowl and coots (Table 1). There are 7 days per census period. The day each cell is to be counted within a census period will be posted in the MERP lab. Counts are to be made in the morning. Census of the selected cells should be completed by 0800 each day. The cells to be counted in any 1 day will be selected so they are adjacent with a tower between them (except for cells 11 and 12).11 is important not to drive on the south dike prior to census as the disturbance will scare birds from the cells. Two observers are required to conduct a census. During a census, 1 observer will walk or canoe through the cells while the second observer is stationed in the The 2 observers will observation tower. communicate by walkie-talkie. In the fIeld, data is to be recorded on data sheet 34. If it is raining, the bird count should be

""NOTE: Always try to minimize the amount of handling of an animal. Needless noise and movement only increases the amount of stress an animal must endure. This can lead to abortions of young in pregnant females caught late in spring. II is often possible to read a recaptured animal's eartags while it is still in the trap or in the handling cone. Simply take 2 straws or small sticks and use these to manipulate the ear of the animal so that the tag can be read while the animal is still in the trap. If done quickly and quietly, this can reduce total handling time as well as stress. This approach also minimizes the risk of losing an animal while it is being transferred to the handling cone. 5.

Introduction

Recording Data (Data Sheet 33)

Record the day as the trapping occasion and the trap number in which the animal was caught. When recording recaptures do not leave blank spaces for 41

postponed 1 day. The effect of rain on walkie-talkies, binoculars, and data sheets will not result in an accurate count. a.

Waterfowl (except Ruddy ducks) To begin a census, park at the north end of the dike on which the tower is located. The count will begin when the tower observer leaves the vehicle and begins to walk towards tbe tower. While approaching the tower, this observer must record all ducks flusbed from the 2 cells to be counted. Only record those birds tbat actually leave tbe cells. Any birds tbat arrive in the cells following tbe start of tbe count should be omitted from final count. Once in the tower, tbe tower observer should ftrst complete tbe blackbird, coot and ruddy duck counts (see below), and then signal the second observer to leave tbe vehicle and begin walking or canoeing through the cells. At tbe start signal, the observer who will walk or canoe through tbe cells sbould enter 1 of the cells and work from north to south in the first cell and then south to north in the second cell. If the cell is flooded, he/she sbould canoe through all the open water and check the emergent vegetation beds. This observer is responsible for identifying tbe ducks as tbey flush. The observer in the tower wiD record data and note any birds that may have been recounted after being flushed once or if flushed birds land in the cell to be counted later that day (so tbey can be subtracted later). In cells with low water, the walking observer must cover the worklanes indicated on the count schedule sheet in the MERP lab. During a count all birds within the boundaries of the dikes, including the ditch, should be counted. The observer in tbe tower should watcb the person in the cell so they can aid in identification as well as identify and count any ducks tbat flush out of sight of the person in the cell. On the field data sbeets (data sheet 34) record pairs, lone males, lone females and others. Lone males are drakes that are not in tbe company of any other birds. If a lone male and a lone hen of the same species occur on 1 cell tbey should be combined and recorded as a pair. A male or males in tbe company of a pair should be recorded in the 'otber' column. A female with a brood sbould be recorded in tbe lone female column and tbe number in the brood in the 'other' column. After 30 June pair use is no longer required, so record only total numbers of each species.

Blackbirds Count only territorial males. Record data for red-winged and yellow-headed blackbirds only. The observer in tbe tower wiD make a complete count immediately prior to the duck count on that cell. A second count on the same cell should be made immediately following the duck count. Record the highest number of birds from both counts for eacb species on the computer.

Coots and Ruddy Ducks The observer in the tower wiD make a complete count immediately prior to the duck count on that cell. The observer canoeing or walking through the cell will make a second count of the coots and ruddy ducks (see data sheet 34). For coots record only total numbers (i.e. do not attempt to determine pairs or sex). Young cools should be included in the total number of coots. On the computer record the highest total (tower and walking count) for both cools and ruddy ducks. Record the ruddy duck data with the waterfowl data.

Nest Counts

Boyd and King (1959) - 'a nest count IS ID theory the best measure of a breeding population: Two nest counts wiD be made each year as follows: Count # 1 2

Beginning Date 17 May 7 June

Four randomly selected zones within each cell will be searched for nests. Within each of the selected zones, 2 observers will make 1 pass with a 20 m nest drag. One observer will walk on the worklane within the zone and the second observer will walk 20 m north or south of the worklane. If a worklane is less than 20 m from the zone boundary, search on the side away from tbe zone boundary. Search for nests between sunrise and noon only. Record all active waterfowl and coot nests between the 2 observers. The observers will also scan an area 5 m on eacb side of their path for active blackbird nests (i.e. a 10 m wide transect for each observer). At each nest site record in a field book, tbe species of bird, zone number, cell number, count period and date. On return to the Station, transcribe tbe data to data forms (see data sbeet 35).

Literature Cited and Additional Readings

MACABE, T.R. 1983. Muskrat population levels and vegetation utilization: a basis for an index. Ph.D. Thesis, Utah State Univ., Logan, UT. 110pp.

ARNASON,A.N. and L. BANIUK. 1978. POPAN2, a data maintenance and analysis system for recapture data. Charles Babbage Res. Cent., St. Pierre, MB. 269pp.

MATHIAK, HA 1966. Muskrat population studies at Horicon Marsh. Wisconsin Conserv. Dept. Tech. BuU. 36. 56pp.

BELLROSE, F.C. and J.B. LOW. 1943. The influence of flood and low water levels on the muskrat population of the Illinois River Valley. J. Mammal. 24:173-188.

MURKIN, H.R 1979. Response by waterfowl and blackbirds to an experimentally manipulated cattail marsh. M.S. Thesis, McGill Univ., Montreal, PQ. 97pp.

BOYD, H. and B. KING. 1959. A breeding population of the mallard. The Wildfowl Trust Ann. Rep. 11:137-143.

MURKIN, H.R., and JA KADLEC. 1986. Relationships between waterfowl and macroinvertebrate densities in a northern prairie marsh. 1. Wildl. Manage. 50:212-217.

CLAY, R.T. and W.R. CLARK. 1985. Demography of muskrats on the Upper Mississippi River. J. Wildl. Manage. 49:883-890. CLARK, W.R. 1987. Effects of harvest on annual J. Wildl. Manage. survival of muskrats. 51:265-272.

MURKIN, H.R., RM. KAMINSKI, and R.D. TITMAN. 1982. Responses by dabbling ducks and aquatic invertebrates to 3n experimentally manipulated cattail marsh. Can. J. Zool. 60:2324-2332.

CLARK, W.R and R.D. ANDREWS. 1982. Review of population census techniques applied in furbearer management. Pages 11-22 in Sanderson, G.c., editor. Proc. Midwest Forbearer Symp., North Central Sec. The Wildlife Society, Wichita, KS.

MURKIN, H.R and B.D.J. BATT. 1987. The interactions of vertebrates and invertebrates in peatlands and marshes. Mem. Entomo!' Soc. Can. 140:15-30. OTIS, D.L., K.P. BURNHAM, G.C. WHITE, and D.R. ANDERSON. 1978. Statistical inference from capture data on closed animal populations. Wildl Monogr. 62. 135pp.

DOZIER, H.L. 1948. Estimating muskrat populations by house counts. Trans. N. Am. Wildl. Conf. 13:372-392.

PALMISANO, A,W., lR 1972. The distribution and abundance of muskrats (Ondatra zibethicus) in relation to vegetation types in Louisiana coastal marshes. Proc. Southeast. Assoc. Game and Fish Commissioners. 26:160-177.

ERRINGTON, P.L. 1%3. Muskrat populations. Iowa State Univ. Press, Ames, IA. 665pp. KAMINSKI, RM. and H.H. PRINCE. 1981. Dabbling duck and macroinvertebrate responses to manipulated wetland habitats. J. Wildl. Manage. 45:1-15.

PERRY, H.R., JR. 1982. Muskrats. Pages 282-325 in Chapman, JA. and GA. Feldhammer, editors. Wild mammals of North America. Johns Hopkins Univ. Press, Baltimore, MD.

KROEKER, D.W. 1988. Population dynamics of muskrats in managed marshes at Delta, Manitoba. M.S. Thesis, Iowa State Univ., Ames, IA. 45pp.

POLLOCK, K.H. 1982. A capture-recapture design robust to unequal probability of capture. J. Wildl. Manage. 46:752-757.

LARSON, 1.S. and R.D. TABER. 1980. Criteria of sex and age. Pages 143-202 in Schemnitz, S.D ., editor. Wildlife Management Techniques Manual. Fourth Edition. The Wildlife Society, Washington, D.C. 686pp.

PROULX, G., and B.M.L. BUCKlAND. 1986. Productivity and mortality rates of southern Ontario pond- and stream-dwelling muskrat, Ondatrazibethicus, populations. Can. Field-Nat. 100:378-380.

LAY, D.W. and T. O'NEIL. 1942. Muskrats on the Texas coast. J. Wild!. Manage. 6:301-312. 43

SEBER, GA.F. 1982. The estimation of animal abundaoce and related parameters. 2nd edition. Chas. Griffm and Co., London, U. K. 506pp.

WELLER, M.W. and C.E. SPATCHER. 1965. Role of habitat in the distribution aod abundance of marsh birds. Spec. Rep. Iowa Agric. Home Econ. Exp. Stn. 43:1-31.

WELLER, M.W. 1981. Freshwater marshes: ecology and wildlife maoagement. Univ. Minnesota Press, Minneapolis, MN. 146pp.

WELLER, M.w. and L.H. FREDRICKSON. 1974. Avian ecology of a managed glacial marsh. The Living Bird 12:269-291.

Table 1. AVIAN CENSUS PERIODS DATE

1 May - 7 May 8 - 14 May 15 - 21 May 22 - 28 May 29 May - 4 June 5 - 11 June 12 - 18 June 19 - 25 June 26 June - 2 July 3 - 9 July 10 - 16 July 17 - 23 July 24 - 30 July 31 July - 6 August 7 - 13 August 14 - 20 August 21 - 27 August 28 August - 3 September 4 - 10 September 11 - 17 September 18 - 24 September 25 September - 1 October 2 - 8 October 9 - 15 October 16 - 22 October 23 - 29 October

Census Period

1 2 3 4 5

6 7

8 9 10 11 12

13 14 15 16 17 18 19 20 21 22 23 24 25 26

Appendix 1

DATA SHEETS 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35.

Weather and Hydrology Data Bulk Density and Percent Organic Matter Physical Environment: Determination of Suspended Solids Gravity Flow Readings Groundwater Levels: Adjacent Groundwater Wells Hydraulic Conductivity Water Chemistry: Delta Lab Sheet Freshwater Institute Lab Analysis Sheet Light Readings For Extinction Coefficient Calculations Solar Monitoring - Light Recordings (E/m'/h) Metaphyton Epiphytic Community Surface Area Calculations % Cover Metaphyton Chlorophyll a Determination Algal Primary Productivity Calculations Inorganic Carbon and pH Determinations Specific Productivity Selection of Macrophyte Belowground Sample Sites Selection of Coring Sites - Belowground Macrophyte Samples Macropbyte Sampling Schedule Belowground Macrophyte Sampling - Lab Sheet Aboveground Macrophyte Sampling - Lab Sheet Turn-Over Plot Field Sheets Fresh Weight/Dry Weight Ratios Root Decomposition Study (1988) - Dominant Vegetation Matching Stems (Permanent Clips) Clip Quadrats Activity Trap Lab Sheet Artificial Substrates Emergence Trap Lab Sheet Invertebrate Dry-Weight Determination Muskrat Trapping Data Sheet Avian Census Field Data Sheet Nest Counts

1 .

WEATHER AND HYDROLOGY DATA Date: a) Weather Data North

Observers: b) Marsh Evap Pans South Cell 11 Cell 12

Time water Added (in) Water Removed (in) Rain

(1mI)

Anemometer Water temp: max min Air temp:

max min

Elevated Anemometer _ __ Record II c) Staff Gauge and P-um-p~Me~ter Readings Cell Time Staff Gauge Pump Meter

Pump

Direction

on off

in out

on off

in out

on off

in out

on off

in out

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on off

in out

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Main Harsh Record II d) Stevens Recorder Cell 6

Cell 46

BULK DENSITY AND PERCENT ORGANIC MATTER

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FRESHWATER INSTITUTE LAB ANALYSIS SHEET

WATER SAMPLE SOURCE DESCRIPTION pH

Site

Code 1

2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21

22 23 24

: Analyt. No. (For lab use only):

Condo

Indicate requested analyses with deviants from fresh water. Requested

Bottle No. (For lab use only)

Date: _ _ _ __

please identify known contaminants or

Analysis Nitrate - N Nitrite - N Ammonia - N Suspended C and N Total Dissolved N Soluble Reactive p Suspended P Total Dissolved P Dissolved Inorganic C Dissolved Organic C Chlorophyll Soluble Reactive Si Chloride & Sulphate Total Suspended Solids Total Suspended Iron CondyctiVity Sodiym potassium Calcium Magnesium Iron Manganese pH Alkalinity Organic acids 49

COI!I!lents

Stored

LIGHT READINGS FOR EXTINCTION COEFFICIENT CALCULATIONS

Date: Time:

Cell : Lane:

COI11Il9nts: LIGHT INTENSITY ABOVE YEG.

WATER SURFACE

1 O.

Date:

5 em

10 em

(~E/m2/S)

15 em

20 em

SOLAR MONITORING - LIGHT RECORDINGS (E/m 2/h)

Date:

12:00-1 :OOam: 1:00-2:00am:

12: 00-1: OOam: 1:00-2:00am:

11:00-12:00om:

11:00-12:00am:

25 em

1 1 .

METAPHYTON Site: Cell : Date: Lane: Water Depth: Distance: Temperature: Time: Vegetation Density (0-5) : Mat Depth': : P : PP : T : PT I

pSy

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¥ METAPHYTON : DENSITY OF SUBMERGED VEG (O,L ,M,H) AND TYPE : COMMENTS:

LIGHT EXTINCTION MEASUREMENTS LOCATION

NORTH

SOUTH

EAST

WEST

ABOVE VEG WATER SURFACE UNDER MAT2 5 CM

10 CM 15 CM

80 CM

'For Eoipelon and Phytoplankton replace mat depth with bottle • . 2For Epjpelon and Phytoplankton delete under mat.

1 2 .

EPIPHYTIC COMMUNITY CELL II: LANE II: DISTANCE: TIME: TEMPERATURE:

SITE : DATE: WATER DEPTH: PERM. QUAD. DEPTH: VEGETATION DENSITY (0-5):

DSv

DSf

DSm

%METAPHYTON: DENSITY OF SUBMERGED VEG (O,L,M,H ) AND TYPE: COMMENTS:

NORTH

SOUTH

EAST

ABOVE VEG

ROD 96~_-7

93"'---_-7 90~_-7 S7~_-7

LIGHT EXTINCTION MEASUREMENTS LOCATION

WEST

S4~_---+

Sl~_-7 7S~_-7 75~_-7 72~_-7

WATER SURFACE

69~_-7

66"'---_-7 63"'---_-7

5 CM

60~_-7

10 CM

57~_-7

51~:--~

15 CM

4S:7 45 : - -7 42:7 39 : - -7 36:

20 CM 25 CM

33~:--'"

30 CM

30:7

27 : - -7 24:. L . - _.......

35 eM 40

21~_-7 lS~_-7

45 eM

15-!-_....... 12~_-7 9~_-7 6~_....

50 eM 55 CM

3.~_....

0"--_....

60 eM

65 eM 70 eM

75 CM

SO eM 52

1 3.

Pg _ of

SURFACE AREA CALCULATIONS

PERMANENT QUADRAT #: DATE SAMPLED: WATER DEPTH SAMPLED:

SITE NAME: CELL: LANE:

Standing Vegetation : #/m2 : : In Qyad.:

Species (Code f)

Total S.A. per Soecies

d=diameter, w=width, l=length, S.A.=surface area

14. ~

COVER METAPHYTON

Cell #: Lane #:

Date:

Mudflat Area

:10

Q4 :W . O.:V.O.:O . Sub.: Macrophyte Coyer:

,,:20,,

* = Possible

:200

Sampling Sites M = distance in meters, W.O. = water depth, V.D. = vegetation density, D.Sub. = density of submerged vegetation, Q = quadrat COMMENTS:

1 5.

CHLOROPHYLL a DETERMINATIONS COMMUNITY: DATE EXTRACTED: AREA/VOL SAMPLED: VOL 90% METHANOL USED: DATA FILE NAME : TIMES OF VORTEXING: 1. 2. COMMENTS : 3.

ABSORBANCE (nm) : Before Acidification:After Acidification I I

No.

Sample Name

Sample Date

665

750

665

750

16.

ALGAL PRIMARY PRODUCTIVITY CALCULATIONS Community/Incubation Date: Comments :

Date Counted:

DPM - Uncorrected (8)=position number L1= L2= : L3=

Sample -a -b -c Dark Control =

( ( ( (

DPM(S)

) ) ) )

: : : :

( ( ( (

: : : ):

( ( ( (

) ) )

I I

: : 1: ):

L4= ( ( ( (

) )

(DPM - Dark Control)

-a -b -c C'4 STANDARDS (X) [ DPM(T)]= INCUBATION TIME-HOURS (T)=

DIC (~g C/25ml)= AREA (VOL)= Productivity Rate: ~g

C fixed/area(vol) per h = DPM(S) x DIC x 1.05 DPM(T) x A(V) x T

Rep] jcates:

a b

-X= SD= SEM= %SEM= where Ll-L4 = ight levels in incubator

) ) ) )

17.

INORGANIC CARBON AND pH DETERMINATIONS DATE ALGAL SAMPLE COLLECTED: DATE WATER SAMPLE COLLECTED: DATE OF INCUBATION: COMMUNITY SAMPLED: NORMALITY OF ACID TITRATED: TEMPERATURE OF WATER SAMPLE: COMMENTS:

SITE NUMBER

TIME: TIME:

H2 S0 4 Titrated (m))

Inorganic Carbon (ug C/25 m))

1 B.

SPECIFIC PRODUCTIVITY COMMUNITY: COMMENTS: SAMPLE SITE

SAMPLING DATE: TEMPERATURE: LIGHT

(uE/m 2 Is2

PRODUCTIVITY (ug C/Lperh2

CHLOROPHYLL (yg Chl alJ)

SPECIFIC PRODUCTIVITY

19.

SELECTION OF MACROPHYTE BELOWGROUND SAMPLE SITES TRANSECT (2 random II's)

SAMPLE SITE NUMBER (locations of sample site in meters from east of transect) 1

)

(

= _m

)

= _m

8 x

= _m

)

9 x

= _m

10 x

= _m

)

11 x

= _m

12 x

= _m

)

13 x

= _m

14 x

= _m

)

15 x

= _m

16 x

= _m

)

= _m

18 x

= _m

)

19 x

= _m

20 --- x

= _m

9 (

10 (

EAST

WEST

- lower of two numbers selected 2_

length of the transect in meters Cell Cell Cell Cell Cell Cell

1 = 155m 2 = 190m 3 = 130m 4 = 135m 5 = 135m 6 = 135m

Cell Cell Cell Cell Cell Cell

= _m

8 (

= _m

5 x

7 (

2 x

)

6 (

4 x

5 (

= _m = _m

4 (

3 x

3 (

)

2 (

Year

Cell

7 = 8 = 9 = 10= 11= 12=

161m 153m 151m 200m 150m 150m

higher of the two random numbers selected 56

20.

SELECTION OF CORING SITES - BELOWGROUND MACROPHYTE SAMPLES CELL SAMPLE SITE

YEAR SPRING

FALL

2 I I

20 For each sample site randomly select 6 numbers between 1 and 10. Do not repeat numbers

21.

MACROPHYTE SAMPLING SCHEDULE Cell Cell

1 & 2

Transects 3&4 5&6

7 &8

9 & 10

2 I I

12 Random number check-off (as the numbers are selected from the random numbers table check them off below to avoid duplication). 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

22.

BELOWGROUND MACROPHYTE SAMPLING Lab Sheet

Date: Cell : Transect:

Cell Staff Gauge Reading (on above date): SAMPLE NO.

VEGETATION AT SITE

VEGETATION : DATE : DATE: WEIGHT OF IN SAMPLE !COLLECTED!SORTEP!RQQTS & RHIZ.

GROUND SAMPLE.

WATER DEPTH

23.

ABOVEGROUND MACROPHYTE SAMPLING Lab Sheet Cell Staff Gauge Reading (on above date): SAMPLE NO.

VEGETATION IN SAMPLE

WEIGHT OF VEGETATION

WATER DEPTH

Date: Cell : Transect:

DATE: DATE COLLECTEP! SORTEP

GROUND SAMPLE.

24.

TURN-OVER PLOT FIELD SHEETS Date:

Crew:

Species:

Cell :

: NEW WATER: GREEN DEPTH: (ORANGE) : SUE II (CM) *F: NF I

I I I I

OLD GREEN *F : NF

GREEN NEW TOPPLED

NEW DEAD (BLUE) *F: NF

I I

OLD

DEAD (BLUE) *F: NF

I I I I

BLUE : TOPPL. : 1988: *F: NF: PSL

Wejght (gms)

Groynd Sample II

yes no yes no yes no

Rhizomes collected: 50 flowering stems: 50 non-flowering stems:

25.

FRESH WEIGHT/DRY WEIGHT RATIOS SAMPLE

FRESH WEIGHT (± .01g)

DRY WEIGHT (± .01g) 80 0 C

26.

ROOT DECOMPOSITION STUDY (1988) - DOMINANT VEGETATION Dominants determined from the 1987 growing season species:~__~~______~~__~_ Collected April 1988, Weights in hundredths of grams.

: :Litter: :Litter: Litter :Ground: : :Site: Bag : Date : Bag : Bag & I O.D . W. at: Date :Sample: :Cell:No. :Number:Deployed:Weight:Ljtter(F.W.>:Collection:Co]]ected:Number: I

•

I !

! .

I ,

I !

27.

MATCHING STEMS (PERMANENT CLIPS) Species:

Date Initiated:

, ,,,

, Site : Cell: Number

,,, ,

, Date Color 1 : Code 2 : Collected

Water Depth

, Ground Litter Weight : Sample (Q.D.W.±0.019l: Number

1Color: B=blue, Y=yellow, P=pink, O=orange 2Code: F=fallen in water, M=missing, C=continuing, I=initiated 28.

CLIP QUADRATS Cell:

Water Depth (±0.5 cm):

(1-10)

Date Collected:

Site Number: Plant Condition , (AD.AL.FU 1 :

Species

Litter Weight «O.D.W. ± 0.019)

Ground Sample Number

1AD=aerial dead, AL=aerial live, FL=flooded live

29.

ACTIVITY TRAP LAB SHEET WD - Water Depth (cm) MAX - Maximum Water Temp. (DC) MIN - Minimum Water Temp. (DC)

Sampling Period:

Date:

WD = MAX = MIN =

30.

ARTIFICIAL SUBSTRATES WD - Water Depth (em)

Sampl ing Period

Date:

WD =

31.

EMERGENCE TRAP LAB SHEET WD WT SST ST

Water Depth (em) Water Temp. (aC) Substrate Surface Temp. (aC) Substrate Temp. (aC)

Sampling Period:

Date:

---

WD = WI = SST = ST =

WD WT SST ST

= = = =

32.

INVERTEBRATE DRY-WEIGHT DETERMINATIONS Date Collected Cell TAXA

No. of Individyals

pan Wt. (gms)

Wt. of Pan: Wt. of & IDverts Inverts

I I

Wt.!lndjvjd. :

33.

MUSKRAT TRAPPING DATA SHEET D

: C: L

a :e'e t

:1 f

, e : 1 t g:h :Y:M:P: @:t ,

:R:Weight: Body :Sex:Age:Occasion Location Comments* Hind (gms):Length'1=F'1=J: Foot (rnm) :, (rnm) 2=M 2=A:

T: i T:e: a:g a:_!

9:e: @:d: ,, ,

,, ,

*the Location and Comments columns should be repeated 5 more times

35.

NEST COUNTS Pate

Cell

Count'

Soecjes 2

'Count Number: 1 or 2 2Species: AOU number

@ Nests

34.

AVIAN CENSUS FIELD DATA SHEETS Observers:

Date: Cell : A. BLACKBIRDS

Red-winged Yellow-headed B. WATERFOWL (except Ruddy ducks) Species

C. COOTS

Pairs

From Tower

Other

Walking Count

Pairs

D. RUDDY DUCKS Tower Coynt Wal king Count 63

Other