SCALER methods Jan 9 2013 v 1.0
Scale Consumers and Lotic Ecosystem Rates: SCALER
Methods Manual
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SCALER methods Jan 9 2013 v 1.0
Contents
1.INTRODUCTIONError! Bookmark not defined.
2.SITE SELECTION AT THE NETWORK, REACH, AND HABITAT SCALEError! Bookmark not
defined.
2.1.Network scale (i.e., synoptic sampling)Error! Bookmark not defined.
2.2.Reach-scale experimental sitesError! Bookmark not defined.
2.3.Habitat scaleError! Bookmark not defined.
2.4.Experimental installation at habitat and reach scaleError! Bookmark not defined.
2.4.1.Reach-scale exclusionError! Bookmark not defined.
2.4.2.Habitat-scale exclusionError! Bookmark not defined.
2.4.3.Basket design and installation at habitat exclosures and experimental reachesError!
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3.Field sampling methodsError! Bookmark not defined.
3.1.Physical characteristics for pre-experimental synoptic site surveyError! Bookmark not defined.
3.1.1.Site locationsError! Bookmark not defined.
3.1.2.Stream width and depthError! Bookmark not defined.
3.1.3.Substrate size distributionError! Bookmark not defined.
3.1.4.Water qualityError! Bookmark not defined.
3.1.5.Canopy coverError! Bookmark not defined.
3.1.6.DischargeError! Bookmark not defined.
3.1.7.SlopeError! Bookmark not defined.
3.2.Consumer survey and voucher specimen at experimental reaches and intensive synoptic sitesError!
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3.3.Whole-stream metabolism of experimental reachesError! Bookmark not defined.
3.4.Whole-stream nutrient uptake and aeration in experimental reachesError! Bookmark not defined.
3.5.Water chemistry sampling of experimental reaches and synoptic sitesError! Bookmark not
defined.
3.6.Habitat delineation of experimental reaches and intensive synoptic sitesError! Bookmark not
defined.
3.7.Whole-stream metabolism at synoptic sitesError! Bookmark not defined.
3.8.Nutrient uptake at synoptic sitesError! Bookmark not defined.
3.9.Algae chlorophyll a and benthic organic matter (BOM) sampling at synoptic sitesError!
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3.10.Metabolism and nutrient uptake of patch exclosuresError! Bookmark not defined.
3.11.Algae chlorophyll a and benthic organic matter sampling of patch exclosures and experimental
reachesError! Bookmark not defined.
3.12.Invertebrate sampling of patch exclosures and experimental reachesError! Bookmark not
defined.
4.Laboratory proceduresError! Bookmark not defined.
4.1.Reaeration SF6 samplesError! Bookmark not defined.
4.2.Streamwater nutrient and ammonium uptake samplesError! Bookmark not defined.
4.3.Benthic organic matter (BOM)Error! Bookmark not defined.
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4.4.Chlorophyll aError! Bookmark not defined.
4.5.Nutrient ratio analysis (C:N:P)Error! Bookmark not defined.
4.5.1.Benthic organic matter C:N:PError! Bookmark not defined.
4.5.2.Biofilm C:N:PError! Bookmark not defined.
4.5.3.Analytical analysisError! Bookmark not defined.
4.6.Invertebrate processingError! Bookmark not defined.
5.Timeline of measurementsError! Bookmark not defined.
5.1.Project timeline of field measurements (among years and across biomes)Error! Bookmark not
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5.2.Experimental timeline (1 year and biome)Error! Bookmark not defined.
6.Data procedures – data managerError! Bookmark not defined.
6.1.Labeling protocolError! Bookmark not defined.
6.2.Field data sheetsError! Bookmark not defined.
6.3.Electronic data sheetsError! Bookmark not defined.
6.3.1.File naming protocolsError! Bookmark not defined.
6.3.2.Raw data entry qa/qcError! Bookmark not defined.
6.4.Backup proceduresError! Bookmark not defined.
7.Modeling proceduresError! Bookmark not defined.
7.1.Data calculationError! Bookmark not defined.
7.2.Workflow storageError! Bookmark not defined.
7.3.Input data file storageError! Bookmark not defined.
7.4.Output data storageError! Bookmark not defined.
8.ReferencesError! Bookmark not defined.
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SCALER- Methods manual
1.
INTRODUCTION
These protocols are for the project SCALER: Scale Consumers and Lotic Ecosystem
Rates. They include site selection criteria, consumer manipulation, measures of ecosystem rates,
and sampling for supporting information. Section 2 describes the reasoning for selection of
specific sites at three different scales: the habitat, the reach, and the network. This section
includes installation instructions for consumer exclosures at the habitat and reach scale. Section 3
details the field methods for metabolism and nutrient uptake to be used at the three scales of this
project as well as additional measures to be taken during the experiment. Section 4 details the lab
protocols for processing of field samples. Section 5 gives a timeline of the field sampling as well
as the experiments with specific tasks to be conducted at each site. Section 6 details data
management and use of field and electronic datasheets while section 7 provides modeling
procedures for metabolism and nutrient uptake. Appendices give material lists overall and by
experimental days, datasheets, electronic datasheets, and other forms.
This document represents a minimal set of measurements to be made as closely as
possible in the same way at each site. Researchers associated with each of the five biomes is
encouraged to undertake additional studies and measurements to answer questions that might be
site-specific or of special interests to the investigators. Coordination among biomes in such
studies is encouraged to find more broadly applicable patterns. As detailed in the authorship
agreement such studies should be detailed to the group.
Parts of this manual were taken from the LINXII protocols led by Pat Mulholland and all
LINXII project participants are thanked for that text.
2.
SITE SELECTION AT THE NETWORK, REACH, AND HABITAT
SCALE
Sampling is conducted at two different scales (reach and habitat) with linkage across the
scales to include the network scale accomplished with modeling. Table 1 describes some of the
models of increasing complexity that we might use to scale metabolism and nutrient uptake from
habitat to reach and from reach to network.
We assess the ability to scale ecosystem rates (e.g., stream metabolism, ammonium
uptake, etc.) and consumer effects on those rates across scales to entire watersheds. Models to
scale metabolism are calibrated using information from habitat-scale measures and the
experimental reaches (3 from each year), and 20 synoptic sites at the network scale that are
studied during baseflow in both years 2013 and 2014. The following biomes take part in the
SCALER study: Luquillo LTER (LUQ), Coweeta LTER (CWT), Konza Prarie LTER (KNZ),
Caribou/Poker Creeks (affiliated with Bonanza Creek LTER; CPC), and the Arctic LTER
(ARC). In the rest of the manual, biome will refer to these five locations. The habitats and
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experimental reaches will span stream sizes ranging from roughly 1 to 500 L s-1 depending on
specific streams within each biome (Table 2).
Within a stream network there will be four types of sites. Habitat experiments will occur
at about 1 m2 spatial area, reach experiments at approximately 10-100 m stream lengths. A series
of 20 synoptic sites will be spread across the stream network, with 6 intensive sites (more
detailed sampling) and 14 extensive (basic sampling).
Data from the extensive synoptic sample sites will be used to inform selection of
representative experimental sites (habitat and reach), if in any way possible as some biomes will
need to select sites based on the reality of physical access. Models will be used to scale from
habitat to reach scales and from reach scales to the watershed. The intensive synoptic survey data
will be used to test and validate model predictions at the network scale. If a model of a particular
level of complexity (Table 1) is able to predict the synoptic metabolism measurements, then we
can have increased confidence that our network scale estimates are reasonable, and that results of
model experiments are useful (e.g., quantifying metabolism of network with and without
consumers).
The whole watershed to be studied will be selected based on the following criteria, while
realizing that optimizing all of these may be difficult: 1) watershed minimally influenced by
humans, 2) watershed large enough to capture important gradients (range of stream sizes
and features such as width or canopy as well as gradient in macro-consumer communities,
and 3) as large a watershed as possible given constraints to be able to accomplish the
experiments and measurements in the largest streams. Following a ‘suitability model’
approach, synoptic and experimental reach selection occurs using a combination of available
LiDAR and GIS data layers of each network within each biome with informed by those with
intimate knowledge of each biome. Criteria for both synoptic and experimental reach selection
are based on ranking of variables such as surficial geology, stream gradient, proximity to access,
and discharge. While stream order will be calculated for networks at each site, discharge will be
used to provide comparative basis between sites with categories small, medium and large (see
Table 2).
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Table 1. Approaches of increasing complexity for reach-to-network scaling of ecosystem rates. In each scenario, estimates of GPP and R (g m-2 d)
from our experimental reaches (n=6, from three different stream sizes) are used as the basis for making a network wide estimate of GPP and R. We
will test which scaling scenario (hypothesis) is most appropriate in each biome, but not necessarily all scenarios will be used for final comparison!
Scaling Scenario
Description
Test
Synoptic Data Needed
Simple Linear
Assume mean/stdev from n=6 intensive sites can be
Ho: Mean/stdev of synoptic sites
GPP and R from synoptic
applied to benthic area of entire network.
are not significantly different from sites.
intensive sites, residuals not a
function of stream order.
Stream-Order
Apply mean from each size class (n=2) to total
H1: Mean/stddev of synoptic sites GPP and R from synoptic
Stratified Linear
benthic area of each order.
of each order are not significantly
sites. Hydraulic
different from intensive sites of
measurements for width vs.
that order
drainage area relationships.
Empirical Model
a) Develop empirical model of GPP and R from
H2: Model predicted GPP and R
Water temp., light, hydraulic
(e.g. GIS and spatial intensive sites as function of temperature, substrate,
consistent with synoptic measured dimensions, substrate, DOC,
modeling approach,
light to benthos, water depth, flow, nutrients (based
GPP and R. (P vs. O)
nutrients, consumers, GPP
such as
on n=6 sites measured over time) and apply to
H2B: Model requires accounting
and R from each synoptic
geographically
network using synoptic relationships and GIS. b)
for the distribution of consumers.
site. Relationships as a
weighted regression) same as (a), but considering consumers.
function of river size.
Process Model (no
Dynamic model of GPP and R that incorporates
H3: Model predicted GPP and R
Same as above plus a greater
consumers)
spatial heterogeneity of drivers (water temperature,
consistent with synoptic
number of stations to
light, DOC, nutrients), C:N:P stoichiometry and
measurements, plus variability
characterize heterogeneity of
effects of upstream transformations (serial processing within the network is greater than
drivers and inputs
and advection). Consumer effects not important.
indicated at synoptic sites alone.
throughout river network.
Process Model (with Same as previous model, but with effects of
H4: Model predictions better fit to Additional synoptic
consumers)
consumers parameterized as a control on processes.
observations than process model
information on consumer
with no consumers.
abundance variability
throughout the network.
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Table 2. Categorized estimated normal water year baseflow discharge values at SCALER study sites.
Synoptic and experimental reach locations attempt to capture the proportional number of small, medium,
and large reaches based on availability within each watershed.
Site Specific Discharge Range (L s-1) by category
Site
Small
Medium
Large
2 - 17
17 - 30
30 - 50
LUQ
2 - 20
20 - 60
60 - 300
CWT
10 - 20
20 - 50
50 - 100
KNZ
20 - 35
35 - 90
90 - 300
CPC
0 - 100
100 - 250
250 - 500
ARC
2.1.
Network scale (i.e., synoptic sampling)
Each year, within each river network, we conduct a synoptic survey of 20 sites (6
intensive, 14 extensive) that will provide base data and validation for various approaches for
scaling reach scale process measurements to entire river networks. The site selection will be
guided by initial modeling, and then accomplished during the initial stages of the first year of
experiment by the postdocs, students, and PI’s at each site. Intensive sites have a more thorough
suite of measurements that would be unfeasible to conduct across the additional 14 extensive
sites. The 14 sites allow better spatial coverage throughout the watershed. The primary
difference between the intensive and extensive sites is that process rates (ammonium uptake and
metabolism) as well as a single-pass characterization of macroconsumers (subset of three
intensive sites per year) are measured at the intensive sites.
Wherever possible, sites are stratified by discharge and other characterizing metrics (e.g.,
stream order), and if necessary by major stream-reach scale variability (Table 2). In an ideal
world, reach heterogeneity is characterized by stream size, but if broad-scale habitat differences
occur (e.g., headwater streams with and without canopy cover; reaches with drastically different
sediment types, etc.), then synoptic sites are sampled to account for this heterogeneity. The
distribution of sites should be weighted by benthic surface area with increasing stream order, and
one at the mouth of the basin. This requires information on average stream width as a function of
stream order. As the metabolic activity and nutrient uptake are driven by benthic surface area,
this approach will allow sampling more weighted to the process of interest rather than simply
relying upon stream length.
Synoptic sites should preferably be unimpacted by water withdrawals and point sources,
and have uniform land use (e.g., little or no agricultural or urban land uses). If impacts are
unavoidable, estimates of water withdrawals, inputs, or point sources should be provided for the
modeling. Some of this information will be available from prior existing datasets. Sites should be
accessible but avoid locations with potential effects on hydrology caused by road crossings and
other impacting factors. If possible, synoptic sites should be at least 10 stream widths
downstream from tributaries or lakes, so mixing is complete. Though junctions may be hotspots
of activity (Benda et al, 2004), determination of this aspect will be left to individual sites if they
are interested in adding such a component. Site selection is first suggested based on models, then
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adjusted for reality by site knowledge of individuals, and as a final reality-adjustment, selection
is refined after field surveys.
Synoptic sampling is conducted at the same 20 sites in 2013 and 2014. We decided to
revisit sites to account for potential interannual variation instead of expanding spatial coverage,
considering that most watersheds will be hard pressed to find 40 sites to study. Repeating the
same sites could also help put varying experimental results in context. The synoptic surveys are
carried out (as much as possible) during the experimental manipulation (40-day period for the
exclosure experiments) and thus the same conditions as the intensive reach-scale measurements
(i.e., similar light regime, flow regime, season, air temperature). If possible, no substantial
changes in discharge should have occurred in the basin in the week prior to sampling, and
discharge conditions should be relatively steady over the course of synoptic sampling. If a
significant storm occurs in the middle of the synoptic sampling, wait until base flow conditions
are achieved, and re-sample a few sites. If the sites are similar conditions prior to the flood, then
not all need to be resampled.
Synoptic surveys include a characterization of the channel geometry (e.g., depth and
slope profiles), bed substrate, in-stream habitat, and adjacent land use. These metrics are
relatively invariant and will only need to be measured once, if possible before the 2013 field
season (i.e., two weeks before experiment starts). Measurements during the experimental period
at all synoptic sites include: water quality parameters (i.e., temperature, dissolved oxygen,
conductivity, turbidity, and pH), velocity, discharge and nutrient chemistry (NO3, NH4, PO4,
DOC, TDN, TDP, PP, PN, PC) (see Table 3). Note, that not all these aspects may not be
analyzed for every site, except for NH4, but we need to take large enough samples to allow for
measurement of these chemical parameters. At the 6 synoptic sites, termed intensive, wholestream metabolism, nutrient uptake, chlorophyll a, and benthic organic matter are measured, and
macro-consumer communities surveyed.
The intensive synoptic survey enables us to: 1) increase sample sizes on some key
metrics above and beyond that collected at experimental sites to obtain sufficient data to develop
and parameterize biological process models at the reach level without huge increases in work
load, 2) provide quantification of the input variables necessary to simulate network-level
metabolism and nutrient uptake, and 3) to test predictions of the network-level simulations at
different locations within the river network (Table 1). Specifically, measurements from a subset
of the intensive sites and information from the experimental reaches is used to develop and
parameterize models of biologically-driven element flux and subsequent metabolism and nutrient
uptake. Once models have been initially constructed (functional formulation and
parameterization) using data from the experimental reaches, network-level models that
incorporate the reach-level biological process models and hydrology will be driven with a subset
(the subset to be determined by the modeling team; see section 7) of the extensive synoptic
measurements. The network-level model predictions will be tested with data not used to
parameterize the reach-level biological process models or drive the network-level models.
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Table 3. Measures to be made at synoptic sites
Measure
Method
Replicates
Comments
Section
All synoptic sites
a) Site characterization survey (2013 only, except if substantial changes between 2013 and 2014; if truly pre-synoptic include width/depth
and discharge listed under b))
Substrate
Transects, pebble
Min. 10 transects, evenly spaced
Done at same time as depth
3.1.3.
characteristics counts
along 1.5x min. reach length, 10
pebbles per transect
Temperature
Transects, probe
Min. 10 transects, evenly spaced
This can help find unexpected areas of
3.1.4.
along 1.5x min. reach length, 3-5
groundwater input
points across the transects
Canopy cover Spherical
Min. 10 transects, evenly spaced
repeat if leaf cover is developing or decreasing
3.1.6.
Densitometer
along 1.5x min. reach length, at
during experiment, or if major changes to 2014
thalweg
Slope
Entire reach,
Use best method available at each site
3.1.7.
clinometer or hose, or
survey equipment
GPS location GPS unit, middle of
3.1.1.
reach
b) During the experiment (2013 and 2014)
Water
Filtered and unfiltered 1 each, at time of synoptic sampling,
~50 ml for nutrient analyses, depends upon
3.5.
chemistry
samples, nutrients
capacity of laboratory and desired level of
cooled on ice and then
replication(note if protocols of individual
frozen
laboratories analyze particulate fraction PC, PN
and PP, then need to filter before freezing and
save filters but no unfiltered samples)
Water quality Conductivity, pH,
Minimum once per site at beginning,
Using probes, snapshot measurements
3.1.4.
turbidity, temperature downstream reach end
Mean width
transects
Min. 10 transects, evenly spaced
3.1.2.
and depth
along 1.5x min. reach length
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Discharge
Flow meter or salt
slug
Minimum once per, constraint point
for flow meter, downstream end for
salt slug
Intensive sites only (n=6, during the experiment)
MacroSampling with
3 sites in 2013 and the other 3 in 2014
consumers
appropriate gear
Metabolism
Single station diurnal
One 24 hour measure per site
change (DO, light)
repeat if base flow substantially increases or
decreases during experiment
3.1.6.
One pass only
3.2.
3.7.
Bott et al.
1996
Needs to be done separately from nutrient release, 2.2.1.
if does not work with travel times, needs to be
Mulholland
done again after pulses
et al. 2001
3.8.
Covino et
al. 2011
3.1.2.
Aeration,
Transient
storage
Nutrient
uptake
Plateau method
1 per site, in conjunction with initial
travel time measurements
Dual station pulse
method
1 per site
Mean width
and depth
Benthic
organic
matter
Chlorophyll a
transects
Minimum of 20 widths and depths
Divide reach into
sections
Multiple samples per section
depending on substrate heterogeneity
weighted by habitat and substrate cover
3.9.
Divide reach into
sections
Multiple samples per section
depending on substrate heterogeneity
weighted by habitat and substrate cover
3.10.
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2.2.
Reach-scale experimental sites
Reaches are chosen so that the major substrata dominating the reach are similar to those
identified in the initial synoptic survey for the specific stream size (see section 2.1.), as well as
the approximate proportion of riffles, runs and pools for each overall reach. At sites where
STREON is planned one of the experimental reaches will be at the STREON site, if possible.
Three experimental sites are
selected to be sampled in 2013 and
three different sites in 2014 (Figure 1).
Sites are chosen to represent a gradient
of stream size with the different stream
sizes of small, medium and large
represented in each year of sampling.
The initial survey of synoptic sites is
used to ensure that experimental reachscale sites are representative of the river
network. Repeated survey of synoptic
Figure 1. Experimental design at the network scale (1 biome)
sites will help control for inter-annual variation.
Reaches are variable in length and set based on the maximum of the shortest distance
required to measure metabolism and nutrient uptake. The shortest length is used because the
consumer removal in long reaches is difficult, particularly in larger streams. The measurement
lengths will depend on travel time and other physical characteristics of a reach site. As control
and treatment reaches within each experimental site are to be compared, they should be
approximately the same length and have roughly the same proportion of pools and riffles. For
specific reach length calculation see section on experimental installation, especially section
2.4.1. and 2.4.2.
2.3.
Habitat scale
Metabolic rates, ammonium uptake, and the influence of consumers are measured for the
dominant substrate types (e.g., cobble or sand) in representative habitats (e.g., riffle and pool).
Survey of substrate type and average width is necessary to set the experimental exclosures and
apportion baskets among habitats in an approximate area-weighted fashion. The basic idea of the
sampling is to capture the dominant types of substrate or cover types at each of the experimental
sites to allow extrapolation from baskets to habitat and from habitat to network. We classify
substrate types via a modified Wentworth scale for inorganic particles and by type for organic
materials as follows (Table 4):
Table 4. Modified Wentworth scale of substrate types
Substrate Type
Median Axis Length (mm)
Cobbles and Boulders
> 64.0
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4.0 – 64.0
0.063 – 4.0
< 0.063
Pebbles and Gravels
Sands and Granules
Silts and Clays
For the purposes of this study, everything greater than cobble size is assumed the same since we
cannot deal with chamber incubations of anything bigger than cobbles. Sand is different from silt
in that we know that flow through sand can create very different biogeochemistry than much
more restricted flow through silt or mud (Dodds et al. 1996).
If the total area of a stream segment being considered is A, and we have n types of each substrate
area cover Sai then:
n
A   Sai
(Eqn 4)
i 1
While we might assume that the proportions of Sai will vary between riffles/runs and pools, this
may not be the case (e.g., when there is a fall/pool structure in steep areas, or when one substrate
type totally dominates a stream). Thus, we need to have decision rules about how we choose
what type of substrate we pick for metabolism and nutrient uptake measurements at the patch
scale as well as what habitat type to place the exclosures in (see section 2.4.3. and 2.4.4.). As we
can at least control for water velocity (riffles and pools) we need to decide if this is more
important or if representing the full range of substrate type is more important (probably in most
cases substrate and water velocity will follow each other so we should be ok). We use this
relationship to find the three most dominant substrates, as defined by the ith term that makes up
80% of the areal cover up to 3 substrate types. A set of decision rules is described in section
2.4.4. to deal with the best way to capture the majority of surface area biological activity.
Apportioning these substrate types by experimental exclosures and in experimental reaches will
be covered in sections 2.4.3 and 2.4.3.
The substrate representation in habitats will be used to select, 1) basket selection in small
scale experimental exclosures, 2) basket selection in experimental reaches, and 3) sampling
strategy for BOM and chlorophyll in synoptic samples.
2.4.
Experimental installation at habitat and reach scale
2.4.1. Calculation of experimental reach length
In general travel times of 30 to 45 minutes are optimal as any shorter, and measurements cannot
be made and any longer then each measurement takes a prohibitively long amount of time. More
importantly, travel time can be used to empirically calculate re-aeration. If possible, prior
estimates of ammonium uptake rates and metabolism rates provide the best way to estimate
experimental reach length, but if not, then from the background ammonium (measured or
estimated) and measured physical variables (travel time and slope) we can calculate minimum
usable reach length.
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To roughly determine reach length, we first need to determine the minimum length to get
a reach-length metabolism measure. Riley and Dodds (in press) determined that for small
streams the equation of Tsivoglou and Neal (1976) gave best estimates of measured aeration
rates using the following equation:
k= C*∆elev./ t
(Eqn 1)
where, C = 0.187 m-1, ∆elev is the change of elevation in meters, t the travel time in seconds and k
the empirically derived re-aeration in units of per second. From this equation we can use the
estimates of Reichert et al. 2009 to estimate minimum practical reach length for a two-station
measurement
∆x = 0.4 v/k
(Eqn 2)
where, v = velocity in m s-1 and k is aeration in s-1.
Using nutrient uptake length data from the LINXI project to calculate the distance it took
to decrease the ammonium concentration the following travel times and reaches need to be
considered to decrease a 5-times background addition by:
30% of that value:
Longest travel time was 30 minutes and longest travel distance was 80 m.
20% of that value:
Longest travel time was 20 minutes and longest travel distance was 52 meter.
If you know your uptake length (Sw) for an individual stream the distance in meters (x) can be
calculated as
x = Sw * (ln (upstream conc.)- ln(desired downstream con.)).
2.4.2. Measurement of travel times combined with aeration and transient storage
An estimate of travel times is conducted before setting each experimental reach site including the
measurement of average width, average depth and discharge. At this point aeration for the
reaches is determined. If time does not permit, travel times can be measured with pulsed releases
before reaches are established and aeration can be done the same day as the nutrient releases
following the pulse release. For pulse measurements of travel time, determined how long it takes
the peak to pass each potential point for a reach (top control, bottom control, top treatment and
bottom treatment reach), starting at the top of the most upstream reach.
Note that this method should also be used at all the intensive synoptic sites, meaning
nutrient uptake first followed by SF6 aeration. Travel time at the extensive synoptic sites is not
needed, as reach lengths are approximates (see section 3.4.1.).
Material list
Injection pump (battery operated, capable of constant 20 mL/min delivery for at least 5 hours)
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Battery
Tubing
Stopwatch
Graduated cylinder, 20 to 100 ml
Meter tape, 50 to 100 m; alternatively an electronic distance measure unit
Injection carboy (20 L) or gas bag, with graduation marks to monitor fluid level
Injection solution (pre-measured amounts of conservative tracers NaCl, NaBr or rhodamine)
Conductivity meter, ion-specific probe, or fluorometer
SF6 gas
For gas sampling:
60-mL syringes (35 per experimental site)
Three-way stopcocks
Evacuate vials for gas (15 samples total)
25 mm filter cartridges (Millipore)
25 mm GFF filters (Zefon)
Sampling bottles (60 mL, 35 per experimental site)
Alternative for gas sampling:
10 ml vacutainers part number
Vacutainer needle holders (BD #364815)
Vacutainer needles (BD# 368607)
Sample bottles for conservative tracer
To be done in advance (night before):
Mix the release solution the night before use: Add conservative tracer to stream water, then fill
the solution into the gasbag. Remove as much air as possible from the bag by squeezing while
holding the septum at the highest point and slightly unscrewing the top. Twist the top closed and
withdraw any remaining air with a syringe through the septum. Add 5mL of SF6 gas per liter of
release solution. Gas is allowed to dissolve overnight.
We use a plateau release for aeration because of concerns about SF6 sample load with a
pulse mass method, concerns about analyzing SF6 concentrations in concentrated stock solutions,
inability to rapidly enough sample for SF6 at the top pulse station, and lack of computational
methods to measure aeration with two pulse stations. Additionally, a plateau allows for
conventional calculation of As/A.
To accomplish the plateau, a continuous release of dye or tracer solution amended with
SF6 is released at the top of the reach (lower treatment reach first due to downstream flow, see
section 2.4.3.). The release point should be chosen far enough above the start of the reach (i.e.,
fence placement) to allow complete mixing. The top of a reach should be placed at a constricted
site to allow good mixing of the pulses for nutrient uptake measurements (as well as easier fence
installation).
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When it is time to start the release, first take a syringe, sample the release solution and
save samples for later analysis of solute concentrations. At this time, also take a background
water chemistry sample. Carefully open the gasbag, press out as much of the SF6 as possible and
attach the tubing allowing as little headspace in the bag as possible. Pull the tubing through the
gasbag holder, attach the tubing to the pump and pump out the rest of the headspace keeping the
gasbag upright. Invert the gasbag and fix holder in a position with the tubing exiting the bab at
the lowest point. Once the tubing is filled, the pump rate is measured. The tube is placed in the
stream at the selected release point and the time noted.
The stream with the added solutes is sampled from the thalweg at the bottom station, the
estimated end point of the reach. The tricky part is how to find the bottom station. Once the
pump is started, go to what you think will be the middle of the reach and wait for the solute to
reach that station. We are looking for about a 30 minute travel time (see section 2.4.1.), so use
the time when the solute starts rising quickly at that station to gauge how far the bottom station
should be, and move to that station.
First take a few baseline samples using the conductivity/ion-specific probe/fluromoeter at
the bottom station. Once the solute hits the bottom station, the measurements need to be made
more frequently (if logging is possible, set 1 min intervals). When the concentration stops
increasing substantially (less than about 1% change per minute) at the bottom station, start
sampling for gas and solute. The travel time to that station is when the concentration has reached
half the plateau concentration, so frequent sampling is needed along the rising edge. Note that
solute concentrations will continue to rise, even for days, as the deeper subsurface zones
gradually come to equilibrium with the water column concentrations, but we will ignore these
very slow processes.
When plateau is reached, sampling can begin. As we are most interested in the aeration
across the entire reach, sampling for gas and inert solutes should be concentrated at the top and
bottom fences of the experimental reaches. Take 5 samples from each of these sites and 3
samples at one or two key points in the reach (e.g., below transition from riffle to pool). At each
location you take gas samples also take replicate samples for the inert solute. This will allow for
calculation of discharge at each of the sampling stations as well as correcting the SF6 samples for
dilution. The dilution will also provide important information for metabolism.
Samples can be collected in vacutainers (VT-6431) or syringes.
For vacutainers, make sure the pressure in the vials is increased so that vials will only fill 50%
full. In the lab, the vials should be equilibrated with atmospheric pressure, and then re-evacuated.
Use a syringe with a three-way stopcock that is at least as large as the vials and insert a needle
into the vial septa. Turn the stopcock valve to allow room air to enter vial. Switch valve to close
off the room, withdraw a volume of gas equal to the vial volume, and withdraw the needle. At
this point the pressure in the vial will be ~50% of atmospheric pressure. The exact pressure is not
critical. After sampling, the volume of solution in vials will be measured to determine the exact
water draw. To sample, insert the vacutainer into the vacutainer needle holders. Be sure to have
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the needle under the water before pushing the vacutainer onto the needle. Record the vacutainer
numbers on the field data sheet with sampling location.
For syringes, use a 60 ml syringe and take 40 mL of stream water per sample. Pull in additional
20 mL of air and allow to equilibrate for a couple hours or shake for 5 minutes. Fill the gas
headspace from the equilibrated sample of the syringe into an evacuated vial either in the field or
back in the laboratory. Remember to label all syringes and vials. Also save samples of the
liquid for measurement of inert solute concentrations.
Once the top station samples are taken, the solute pump can be stopped. Note time. Work down
and sample the downstream sampling stations from top to bottom.
Once all the samples are taken, check the pump rate again and then turn off the pump.
Make sure to note the time. Continue monitoring the downstream station for inert solute until the
concentration is at least within 1.5x of the background concentration. The full shape of this curve
is necessary to calculate transient storage characteristics. The transient storage could be helpful
to explain the inability to scale baskets to reaches, the amount of respiration measured, and some
characteristics of the nutrient pulse releases. Once the inert solute concentration has returned to
background, repeat the plateau procedure for the control reach. If the transient storage is large,
and the return to baseline slow, the treatment reach at a second experimental site can be used (if
a second probe is available). If you are limited on inert sampling meters, you may need to wait.
For intensive synoptic sites there is of course only one plateau to do.
2.4.3.
Reach-scale exclusion
Material list
Fencing (3 or 4 ft high, 4 x 1.5 stream widths long for each experimental site (3x), mesh size see
Table 5, suggested place to order from www.tepinc.com )
Clippers
Shovels
Rebar (~1 every 30 cm for the 12 fences) maybe t-posts if high flow
Zip ties
Mallet or sledgehammer
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SCALER methods Jan 9 2013 v 1.0
For one experimental manipulation, reach-scale consumer manipulations are set up at
three sites across the gradient of stream sizes (see reach-scale selection (2.2.) for further details,
Figure 1). Each site contains three reaches: a) an upstream control reach with consumers
removed and returned, b) an area with site-representative habitats, where small mechanical
consumer exclusion/control remove large consumers around baskets at habitat-scale (see habitatscale exclusion below), and c) a consumer removal reach (Figure 2).
Figure 2. Experimental design at the reach-scale (1 site).
Metal fencing with biome-specific mesh size is used to delineate reaches, and exclude
macroconsumers such as fish, crayfish, shrimp, or salamanders from the downstream treatment
reach. Note, our macroconsumers are defined based on mesh size, and mesh size is chosen to
exclude the dominant large animal in each biome. Macroconsumers are not defined based on
functional group. Mesh size is determined for each biome, but within a watershed mesh size will
remain constant at all experimental sites. The mesh sizes selected for the biomes are described in
Table 5. Mesh screen is placed upstream and downstream of control and treatment reaches.
Constricted locations are chose to make it easier to install fences and to facilitate nutrient uptake
measurements. At the designate upstream and downstream locations a trench (deeper the better)
is dug across the stream. Rebar is hammered into the sediments spaced every 30 cm unless
strong flow requires more support. A piece of fence is cut to stretch the channel and up the
banks, and a 20 cm lip bent at a 90° angle (bending on land allows for a clear angle). The fencing
is then placed across the stream with the lip facing upstream and the vertical part placed against
the rebar. The stream sediments removed during trench digging are then shoveled onto the lip to
bury the fencing. Zip ties are used to attach the fencing to the rebar. A wire with electric current
(e.g., using a fence charger) can be run near the stream bed to inhibit consumers from burrowing
through gravel into the exclosure (if deemed necessary based on site characteristics). If
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SCALER methods Jan 9 2013 v 1.0
particulate organic matter transport and/or flow velocities are high, a second fence with a larger
mesh size can be installed across the stream (without the burial) at the upstream end of the
experimental site to help prevent fences from being blown out by spreading the pressure across
multiple fences. In some cases a row of rebar or PVC pipes without fencing can also be
considered. If multiple fences are installed they should be spaced far enough apart to allow for
cleaning off of organic material. It is recommended that fences are cleaned (scrub brush works
well) as often as possible. This may require daily cleaning at some sites.
Table 5. Mesh size for macro-consumer exclusion at reach- and habitat-scales.
Biome ID Biome description
Selected mesh
size
6 mm
LUQ
Luquillo LTER
CWT
Coweeta Hydrological
Station
6 mm
KNZ
Konza Prairie LTER
3 mm
CPC
Caribou/Poker Creeks
LTER
6 mm
ARC
Arctic LTER
6 mm
18
Reasoning
Consumer relatively large and high
organic matter transport makes
fence maintenance difficult
Particulate organic matter transport
high and flow velocity low, macroconsumers medium
Particulate organic matter transport
and flow velocity low, macroconsumers small
Consumer relatively large and high
organic matter transport makes
fence maintenance difficult
Consumer relatively large and high
organic matter transport as well as
high discharge makes fence
maintenance difficult
SCALER methods Jan 9 2013 v 1.0
2.4.4.
abitat-scale exclusion
Material list
Exclosure wood frame: 19 in long wood pieces of 0.5x0.5in (WxH), screwed together in a square
with a 20.5in piece across the diagonal for stability. 8 per experimental site, 3 experimental sites,
24 20.5” pieces, 96 19” pieces, Size and shape may need to be modified slightly for the
narrowest streams, but we still need to be able to get all 5 baskets into each side of the exclosure
Staple gun and staples
Fencing (3ft high, 50ft long per site, mesh size biome-dependent, see Table 3)
Clippers or heavy duty scissors
Zip ties (~100)
Rebar (8 pieces 4ft long 1/2in diameter)
Mallet or sledgehammer
Shovel
Strawberry baskets (10 x 10 x 6 cm), 90 per site for exclosures
Liners for strawberry baskets if holes are too big to hold native substrate (these should be
synthetic fabric of some sort, could be mosquito netting, no-seeum netting, or wedding veil
material)
Stream gravel (mixed in bucket)
Bucket to collect and mix gravel
To be done in advance (rebuild year 2 if necessary):
Build exclosure frames and line strawberry baskets (if necessary)
Frame construction: cut four 19-in long sections of wood pieces, and one 20.5-in piece.
Drill a hole in one end of 19-in pieces, approximately 0.25-in from the edge. Use wood screws to
fix pieces together to create a square frame. Place the 20.5-in piece along the diagonal and fix in
place using a staple gun at the four corners and both from the top and bottom (Fig. 3a).
Basket lining: If the dominant sediments in a habitat are too small to be held within the
baskets (<1 cm), the baskets will be lined with nylon netting. Netting is placed on the inside and
fixed at the top using zip ties. The point with the mesh is to mimic the natural subsurface flow
rates, too fine and flow will be impeded, too coarse and they will wash out.
To be done during experimental setup:
Construct and install exclosures
Construction: Micro-scale exclusion will be accomplished with mesh fences around 0.48
x 0.48 m frame. A piece of mesh is stapled to the bottom of the wood frame (Fig. 3b) followed
by a length of fence stapled along the diagonal and three outsides. The mesh from the diagonal
and the outside will be joined using zip ties (Fig. 3c).
Installation: In the stream, a 0.5 x 0.5 m plot will be shoveled approximately 10 cm deep
with a corner pointing upstream (i.e., the diagonal is parallel to stream flow). The exclosure is
then placed in the excavated plot. Half the downstream edge will be closed, while the other
downstream-half is open to create a non-exclosure control for a paired design with equivalent
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SCALER methods Jan 9 2013 v 1.0
hydrologic alteration (developed by Murdock et al. 2010, Fig. 3d). Five strawberry baskets filled
with substrate collected from the stream (see below will be placed in both the enclosed and open
side of the exclosure frame (Fig. 3d). The sediment from the excavation will then be placed
around the baskets until the top of the baskets and the sediments are level with the stream bottom
outside the exclosure.
a)
b)
c)
d)
Figure 3.Construction and set up of habitat exclosures: a) diagonal piece stapled to external square; b)
mesh stapled to one side of square; c) Mesh stapled to side of diagonal and around three sides of the
square and diagonal zip-tied to close off one side; and d) exclosure placed in sediment with strawberry
baskets on both sides of the exclosure (flow is from left to right).
2.4.5.
Basket design and installation at habitat exclosures and experimental reaches
Material list
Native substrates, collected from within the reach based on substrate sizes (detailed below)
Strawberry baskets, 80 for exclosures, 32 for sampling experimental reaches, per site per year
Mesh lining (if deemed necessary)
Plastic baskets (10 x 10 x 6 cm) (Strawberry baskets) will be modified to use as experimental
units at the habitat scale and at the reach scale. Substrate used in baskets and basket mesh sizes
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SCALER methods Jan 9 2013 v 1.0
will be roughly consistent with planned STREON protocols and determined from sediment size
structure analyses following the rules below:
1. At sites with streambeds consisting mostly of sand (2 mm) or smaller particles, baskets
shall possess wall and bottom mesh openings of 1 mm and 0.5 mm, respectively.
2. At sites with streambeds consisting of cobble, pebble and gravel, basket will be lined
with finer mesh to contain finer sediments (approximately 20th and 10th percentile of the
particle size distribution). The 200-sample pebble count (Bevenger and King 1995; see
Table 2) from the pre-synoptic sampling is used to quantify the sediment composition to
determine percentiles.
3. In reaches where streambed sediments consisting mostly of boulders and large cobble,
the maximum mesh opening size (for both walls and bottoms) shall be 10 mm. At such
sites, rocks can be picked that will fit into the baskets.
To determine placement of exclosures and substrates used in exclosures the entire
experimental reach site is surveyed for pebble counts and average width as per method for
presynoptic sampling. In addition, it is noted if each transect is a riffle or a pool. Note a run
might be considered a riffle or a pool, it does not matter where you draw that line within a
biome as long as you are relatively consistent within that biome. In this section riffle/run is
referred to as riffle from this point forward. Thus, the transects are separated into riffles and
pools. We are interested in scaling up from area-specific rates in baskets to whole-stream rates
and we thus need the relative area in riffles and pools. The total area in riffles Ar is calculated as
Ar = average width (m) of riffle transects* length (m) between transects* number of
riffle transects.
The total area in pools Ap is calculated the same way. The proportion of area in riffles is
calculated as
Proportion of Riffle = Ar/ (Ar+ Ap)
Then the habitat exclosures are apportioned relative to the relative area of pool and riffle
Table 6. How to determine the total number of exclosures to be placed in each habitat type
Proportion Riffle
0 – 20%
20 – 40%
40 – 60%
60 – 80%
80 – 100%
Riffle habitat
exclosures
0
2
4
6
8
Pool habitat
exclosures
8
6
4
2
0
Within each habitat type exclosure (riffle or pool), we need to apportion the baskets within
exclosures based on relative abundance of substrate types. Sediments used to fill the baskets are
collected in a bucket per substrate type (Table 4; bedrock will be excluded due to size constraints
of baskets) and baskets within one exclosure contain only one substrate size for all five baskets.
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SCALER methods Jan 9 2013 v 1.0
If possible the exclosures are also placed in the native substrate that fills the baskets. If 80% of
the substrate is of type 1 (S1) then all of the exclosure baskets are filled with S1 in that habitat. If
S1 and S2 together make up >80% of the surface cover, then two main options present
themselves (both sizes are approximately equal [left], or one dominates [right]):
Table 7. Exclosure apportionment with two dominate substrate types (S1 and S2)
S1 ≈ S2 >>S3
Habitat
Exclosures
2
4
6
8
S1
1
2
3
4
S1 ≈ 2*S2 >>S3
S2
1
2
3
4
S1
1
3
4
2
S2
1
1
2
3
It is possible that three substrates S1, S2 and S3 make up >80% together, in this case different
relative abundances present themselves:
Table 8 Exclosure apportionment with three dominant substrate types
Habitat
Exclosures
2
4
6
8
Habitat
Exclosures
2
4
6
8
Habitat
Exclosures
2
4
6
S1
1
2
2
3
S1
1
2
2
3
S1
1
2
3
S1 ≈ S2 ≈ S3
S2
1
1
2
3
S1 ≈ S2 ≈ 2*S3
S2
1
1
2
3
S1 ≈ 2*S2 ≈ 2*S3
S2
1
1
2
22
S3
0
1
2
2
S3
0
1
2
2
S3
0
1
1
SCALER methods Jan 9 2013 v 1.0
4
3
1
Each exclosure has 5 baskets buried flush with substrata in both the open and the enclosed
side (n=80 baskets). Additionally, pairs of baskets are buried flush with the ambient substrata in
the stream bottom of pools and riffles. Locations of the baskets and substrates in them in the
control (n=16) and treatment reach (n=16) are given below. Baskets in the exclosures are used
for habitat metabolism and NH4 uptake in chambers (3 per exclosure, section 3.10.). The
remaining two baskets and the pairs of baskets in the control and treatment reaches are used for
invertebrate sampling (1 basket; see section 3.12.) and chlorophyll a and benthic organic matter
(1 basket; see sections 3.11.)
In the experimental reaches, there are 8 pairs of baskets per control and 8 per consumer
removal reach to use for inverts, BOM and chlorophyll. Based on survey of percent riffle and
pool, baskets pairs are apportioned as
Table 9 Apportioning baskets for invertebrate BOM and chlorophyll
Proportion Riffle
0 – 20%
20 – 40%
40 – 60%
60 – 80%
80 – 100%
Riffle Basket
Pairs
0
2
4
6
8
Pool Basket Pairs
8
6
4
2
0
You will notice that the number of pairs equals the number of exclosures. Thus, the tables above can be
used also to apportion pairs of baskets assigned to each substrate.
3. Field sampling methods
3.4. Physical characteristics for pre-experimental synoptic site survey
3.4.1.
Site locations
Material list
GPS
Rangefinder/ Tape Measure
All sample sites are surveyed using a GPS, with points also identified on paper maps, and sent to
the modeling group to ensure accurate spatial location on the river network. Synoptic sites are
assessed prior to the experimental manipulation in 2013. Initial synoptic site surveys consist of
reaches that are 1.5x the minimum-determined reach length of the appropriate experimental
stream size, with a minimum of 10 transects perpendicular to the flow direction of the thalweg
and spaced evenly along the reach. This sampling is also conducted at each experimental site
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SCALER methods Jan 9 2013 v 1.0
before the experiment starts to determine placement of exclosures and baskets (see section 2.4.3).
Every transect will be classified by average width:depth ratio as riffle, or pool. Again, runs and
riffles can be combined, and it is not as important how they are defined as that the definition
remains consistent across sites within each biome. Each transect will have 3-5 evenly spaced
points along each transect depending on the width. At each point along a transect measurement
depth, classification of substrate (e.g. clay, sand, cobble, bedrock, macrophyte) and benthic cover
is completed. Also, temperature surveys are done at each transect to establish any areas of strong
groundwater influence; no reaches with significantly flowing side channels are used. In addition,
water quality and canopy cover will be surveyed at each transect at the thalweg point. Discharge
is measured at a subset of transects while slope is measured over the 200 m reach. If possible
triplicate ammonium samples should be taken at each synoptic site to help guide calculation of
ammonium addition rates for ammonium uptake experiments.
3.4.2.
Stream width and depth
Material list
Rangefinder / Tape Measure
Meter stick
Stream width and depth are surveyed along at least 10 evenly spaced transects of the reach at all
synoptic sites moving in a downstream to upstream direction. Width is measured using an
electronic distance measure or a measuring tape. Water depth is measured at 5 locations
approximately evenly spaced across the stream channel using a meter stick and relative to water
surface. Note water depth is difficult to measure accurately in shallow streams and average
velocity, average width and discharge can be used to more accurately calculate average depth
over a reach.
3.4.3.
Substrate size distribution
Material list
Ruler
At all synoptic sites, substrates will be selected at all transects of the reach. A minimum of 200
particles should be surveyed in the reach, meaning the number of substrates per transect depends
on the number of transects chosen (min. 10). The median axis of each particle will be determined
using a gravelometer or a ruler.
We classify substrate types via a Wentworth scale modified for our project to simplify
sampling, for inorganic particles and by type for organic materials as follows:
Table 10 modified Wentworth scale for this proposal
Substrate Type
Median Axis Length (mm)
Cobbles and Boulders
> 64.0
Pebbles and Gravels
4.0 – 64.0
Sands and Granules
0.063 – 4.0
Silts and Clays
< 0.063
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We will also note cover at sample point by wood, leaf packs, dense algal growths, bryophytes or
other macrophytes as these could dominate the biological activity and cover the surface area.
3.4.4.
Water quality
Material list
Multiparameter sonde (use YSI ProODO for temperature)
At all 20 synoptic sites general physical parameters are measured at the bottom of each site (pH,
conductivity, turbidity), except for temperature. Temperature is surveyed every 5 m transect
along a 200 m reach at five evenly spaced locations across the stream to detect potential
groundwater influences. All metrics are measured during the initial survey of synoptic sites. At
extensive sites, they need not be measured again unless there have been major changes (e.g.
floods, continued drying during the experiment). At the intensive synoptic sites they should be
measured again when ecosystem rates are measured.
3.4.5.
Canopy cover
Material list
Spherical densiometer
Canopy cover is surveyed at all experimental and synoptic sites. A spherical densiometer
(http://www.cspforestry.com/Spherical_Crown_Densiometer_p/densiometer.htm) is used to
estimate canopy cover in the middle of the stream channel at all transects along the reach. The
protocol is printed on the densiometer, and is as follows: Measurements are made in each
cardinal direction (i.e. facing north, east, south, and west) and make a note of the direction of
channel from upstream to downstream at the point of measurement (i.e cardinal direction of
water flow downstream at that point). Hold the densitometer in the same location, and move your
body around it when changing positions. To take readings, hold the instrument level, 12" to 18"
in front of body and at elbow height. Ensure the bubble level is centered. Assume four equispaced dots in each square of the grid and systematically count the dots equivalent to open
quarter-squares. All sites should follow the protocol regardless of species height and
composition, but acknowledge in sites with low shrubs (<1 m) there may be an underestimation
of cover.
Calculate percent cover according to the densiometer’s manual: Multiply the count of open
quarter-squares by 104 to obtain percent of overhead area not occupied by canopy. The
difference to a hundred is an estimation of percent overstory canopy.
3.4.6.
Discharge
Material list
Measuring tape
Flow meter
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SCALER methods Jan 9 2013 v 1.0
Pulse release method
Salt or dye solution
Conductivity meter or fluorometer
Sample bottles (at least 20)
Plateau method
Metering pump and tubing
Battery
Graduate cylinder
stopwatch
Salt or dye solution (can be mixed quantitatively in field with stream water)
Conductivity meter or fluorometer
Sample bottles (at least 7)
Discharge can be measured with inert solute releases or velocity meters. For shallow streams, inert solute
releases are preferred. There is a tradeoff between salt slugs and plateau methods for discharge. Detailed
discussion of both methods can be found at http://pubs.usgs.gov/twri/twri3-a16/pdf/TWRI_3A16.pdf. Measurement of Discharge Using Tracers U.S. Geological Survey, Techniques of WaterResources Investigations, Book 3, Chapter A16 By Frederick A. Kilpatrick and Ernest D. Cobb. This
source should be used as our method reference. The source also contains the equations and calculations
needed to calculate discharge as well as preparation of stocks and how much needs to be used.
Both pulse and plateau methods take the same amount of time, because the plateau is reached in the same
amount of time as it takes the pulse to completely travel past a downstream point. Pulse method does not
require a metered pump, uses less mass of solute and release solution does not need to be mixed
volumetrically. Sample burden is much greater with the pulse method as samples need to be taken and
analyzed for every part of the pulse to accurately calculate the discharge.
For flow meter method, Discharge is measured at a constrained location of the stream using a
flow meter. At a minimum of 15 points across the channel flow velocity is measured at 0.6x
depth, and flow, distance across the channel and depth recorded. Alternatively, a salt slug is
released and time until the peak reaches the downstream end (as measured by conductivity), is
considered travel time. The dilution of the salt solution in turn is discharge.
3.4.7. Slope
Material list
Clinometer, hand level and pocket rod or, a long hose
If available, a handheld level with a 5x magnification and a stadia rod can be used with standard
surveying techniques.
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SCALER methods Jan 9 2013 v 1.0
Slope can also be measured using a clinometer from the upstream to downstream end of the 200
m reach. One person stands at the upstream end whit the other person at the downstream end.
The clinometer is then aimed from one person to the other at the same height.
Alternatively, especially when line of site is obscured, get a long hose. Use a tape to get
exact stream length between two points (distance along the run). Lay hose in water and fill.
Upstream person puts opening to hose right at surface but able to refill. Person at downstream
lifts hose till pressure head allows no more water to flow out of the opening. The distance
between said height and the surface of the water is the rise. Slope is rise divided by run distance.
Note, most sites do not have accurate enough GIS and LIDAR to get good slopes over the
length of experimental reaches. However, if they do, that method is also acceptable.
2.6.
Consumer survey and voucher specimen at experimental reaches and
intensive synoptic sites
Material list
Note book, pencil
Backpack electroshocker
Dip nets
Buckets
Aerators/bubblers
Measuring boards
Seine
Minnow traps and bait
Waders
Nalgene bottles (for voucher specimen)
Formalin
Label paper
Consumer surveys are conducted in control and treatment reaches, as well as in the reach
containing the habitat exclosures. Appropriate sampling method(s) are selected for each biome
and reach site. The goal is to remove as many consumers (species and individuals) as possible,
and each biome needs to determine the best methods to achieve this goal. The use of multiple
gears (e.g., seines and electrofishing) is often the most effective way of removing different
species. Independent of gear, control and treatment reaches are sampled in multiple passes for a
depletion sampling. A minimum of four passes with diminishing catch are conducted to allow for
population estimates since the first pass is often not useable for depletion calculation. For each
pass, individuals are identified and length measured if weight-length relationships are available
(nose to fork/fin depending on how relationship was developed). Otherwise, such relationships
should be established. Once a size distribution is established for a species, individuals can be
counted for size classes. The “patch” reach containing the habitat exclosures is only be surveyed
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SCALER methods Jan 9 2013 v 1.0
with one pass, but should be walked through the same number of passes as exclosure reaches to
replicate disturbance. A subset of intensive synoptic sites (three in 2013, three in 2014) is also
sampled with a single pass. Individuals from riffle and pool habitats are to be kept in separate
buckets to allow determination of differences in treatment effects based on habitat type.
Selection of holding containers is made to minimize consumer mortality (e.g., flow through
containers or frequent water exchange).
The treatment reaches are sampled at the beginning and end of the experiment. Additional
sampling at day 11 and 22 of the consumer removal experiment is used to test for exclusion of
large consumers as well as removal of consumers that have outgrown the selected mesh size of
the fence. As patch exclosures were disturbed during installation there should be no consumers
in the closed side, but a quick check with an electroshocker may be warranted. Consumers in the
removal reach are released below the downstream fence (to avoid inflating the number of
consumers in the middle reach where the small exclosures are) or above the top most fence if
consumers have an upstream migratory behavior. The control reaches are sampled at the
beginning and end of the experiment also, but individuals returned into the reach. During day 11
and 22 the same disturbance from the treatment reach will be imparted on the control reach. The
patch reaches are disturbed whenever sampling occurs in the treatment reach and sampled with a
single pass at the end of the experiment to compare densities to other reaches.
Voucher specimens representing large excluded consumer communities in each biome are
collected and deposited at the University of Kansas Museum of Natural History. This is a
necessary step in data management, and is part of the proposal. Several individuals representing
each species collected in each stream and will be fixed in 10% formalin. Specimens are held in
durable and appropriately sized container (e.g., 500 mL Nalgene bottles) and include a
waterproof label that provides the following information: GPS coordinates of the capture
location, waterbody description (e.g., Kings Creek), date of collection, names of collectors, and
field number to reference field notes. Copies of field notes will accompany the specimens and
should include: sampling gear and methods used to collect specimens, collection permit
information, and other information collected at the time specimens were captured (e.g., weather,
water quality).
3.5.
Whole-stream metabolism of experimental reaches
Material list
4 4ft ½ in rebar
Mallet
4 YSI ProODO meters with extended battery compartment
4 Odyssey irradiance meters
Bull’s eye level
Zip ties
NIST traceable thermometer for calibration
Bucket
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erator
Whole-stream metabolism at the reach scale is measured using a two station method.
Dissolved oxygen probes and PAR meters are placed below the upstream and above the
downstream exclosure fence of both the control and the treatment reach outside of potential
influence from the exclosure fences. The DO and PAR meters are calibrated by methods detailed
below. A rebar is hammered into the sediment and the PAR meter zip tied to the top and adjusted
with a level. The DO meter is zip tied to be parallel to stream flow and in the water column
without touching the sediment.
The PAR meters need to be initiated with a computer before departing for the field, the
location is then noted with corresponding logger number once placed. The DO meters are
programmed in the field and should carry the designations specified in section 6.1. Both loggers
will be set to log at 10 minute intervals and remain in the field for 5 days if possible. This will
allow cross comparison of reaches as days are slightly offset by experimental demand.
Reaertion and stream widths are determined at the beginning of deployment during the initial
(Day 1-5) measurements and if the conditions have changed substantially during the experiment
on collection day for the end of the experiment deployment (Day 26-31) using SF6. Methods are
detailed in the whole-stream nutrient uptake section below (3.5.).
Calibration of sensors for reach and chamber measurements:
PAR sensors
 Initially, PAR sensors are all compared to a standard probe calibrated at NEON. This will
be done at the beginning of each year at KSU or NEON.
 Prior to work at each site, all probes should be run simultaneously under the same light
conditions for 20 minutes and then under dark conditions. Apply calibration factors and if
readings vary, return the malfunctioning probe for immediate re-calibration or repair to
the manufacturer (Odyssey Data Recording Systems).
Oxygen probes
 Each year all temperature sensors will be calibrated against a standard probe calibrated at
NEON. This will be done at the beginning of each year at KSU or NEON.
 Temperature sensors on the oxygen meters should be calibrated against a known
standardized thermometer at two points at each site. An ice water bath and room
temperature are standards (http://www.astm.org/Standards/temperature-measurementstandards.html). Fisherbrand* Red-Spirit* General-Purpose Laboratory Thermometers
meet or exceed NIST* and ANSI*/SAMA tolerances for accuracy15-041-5A and do not
have problems with mercury contamination if broken. If a probe is not at the correct
temperature, note how far off it is. If it is more than 0.2 ⁰C off, swap probes with a meter
that is working to determine if the probe or the meter are malfunctioning. Immediately
send the malfunctioning meter or probe for repair.
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


Oxygen probes should be calibrated each day that they are used, and once each hear
against NEON equipment. Any additional probes used at individual sites should be
calibrated against project probes at the beginning of the work at each site each year.
Atmospheric pressure values are calibrated against the closest active weather station.
Barometric pressure can be entered into probes directly. Be certain to run the probes
simultaneously for 30 minutes to ensure that the Barometric pressure readings are similar
across all probes at the end of the run.
Dissolved oxygen: It is absolutely essential that O2 probes be calibrated and ensured to all
be stable. Each day before deployment, all meters are calibrated together in the field (if
this is not possible or outdoor temperatures are above 30°C, then in the lab immediately
before traveling to field site). First, ensure that all the barometric readings are the same, if
not set them to be the same. Then calibrate the sensors in moist air (with the plastic cover
over them with a wet sponge). Be certain that the probes are stabilized for temperature
and at the same temperature for this step. It is best to be certain all probes and meters are
in the shade. After air calibration, all meters are placed in the stream or in a bucket with
stream water and constant mixing from an air pump (the bucket is preferred as we cannot
be certain to have all 20 probes in one place in a stream with exactly the same O2) and
allowed to log for 30 minutes at a 5 minute interval. Temperature is attempted to be kept
constant by placing all probes in the same location in flowing water or in a bucket that is
shaded and place in flowing water. Check the O2 reading, and repeat calibration until all
meters give the same results (within 1% of the mg/L reading) before deployment. All
meters to be deployed at one reach site (n=4), for chamber measurements (n=6), and for
synoptic sites (n=6), are calibrated as a group each time they are used. At the end of
deployment, meters are again placed together at one location (or an aerated bucket if
necessary) for 30 minutes, logging at 5 minute intervals. This allows for correction
assuming a linear drift in calibration over the period of measurement if meters do not
read the same value post-deployment. However, the meters should not be used again until
stability of the meter is determined. To trouble shoot drifting probes, switch probe to
meter known to be working, and making measurements both with the potential problem
meter with good probe and problem probe with good meter. This way you can isolate
which is causing the problem. The problem meter or probe must immediately be repaired
to have equipment ready for the next set of measurements. This means contacting YSI
repair and fedexing the equipment to them.
3.6.
Whole-stream nutrient uptake in experimental reaches
Material list
Meter tape
Open-topped bucket/garbage can (note: the injection volume may depend on how much NaCl is
needed; a larger carboy/release container will allow more NaCl or Rhodamine to be dissolved)
Injection solution (i.e., appropriate amounts of NH4Cl and conservative tracer – Cl or Br)
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Salt slug
Filter apparatus (GFF, Zefon type filters)
Sampling bottles (~50 60mL acid washed Nalgene, pre-labeled, minimum acid washing
requirement, 1 h soak in 0.1 molar HCl followed by at least 6 DI rinses. Sample bottles should
also be rinsed with a small amount of sample water at least once before filling)
Conductance meter, Bromide meter, or Fluorometer
GPS
Field data Sheet(s) (Nutrient_uptake.docx)
Optional: Sampling bottles (250 mL) for collecting unfiltered samples in the field with same
label numbers as syringes for filtering
Cubitainers (~1 L depending on stream size)
The NH4 pulse (slug) addition experiment consists of an instantaneous addition of NH4
(NH4Cl) together with a conservative tracer (e.g. NaCl or NaBr) and sampling of water at
various times during the “breakthrough” of the slug at one location downstream (based the
TASCC method; Covino et al. 2011). The idea is to capture the entire concentration-time curve
for both tracers at the sampling site. The injection solution consists of stream water and the
conservative/active tracers.
To be done in advance:
Carefully calculate the masses of solutes needed, based on a recent discharge
measurement (it is ideal to measure discharge the morning of the experiment, but acceptable to
measure discharge the day before so long as flow did not receive new inputs, if the SF6/
discharge/ travel time measurement is done correctly the day before this should give discharge).
A calculations template is available in the SCALER all dropbox. Weigh out all solutes to 0.1%
accuracy and store them in labeled Ziploc bags. It is most important you know exactly the mass
you added, but it does not need to be the exact mass targeted in your prior calcuations. Note mass
of all solutes in lab book and at appropriate place on field data sheets.
An upstream injection site should be identified. This site should be well-mixed (at a
constriction is best and engineered to push most flow through a narrow area). The engineering
can be done with plywood panels, or natural rocks lined with a plastic. It is probably best to do
the major movement of stream substrates the day before to minimize disruption.
When the nutrient addition experiment occurs at an experimental site, the injection
location should be upstream of that experimental reach upstream net (Fig. 4). We will put the
release site a meter or two above the upper fence. While this may not allow for complete
lateral mixing at the very top of the experimental reach, it will obviate the need for two pulse
sampling stations. The lower sampling station should also be at a constriction.
For the intensive synoptic sites, use discharge, average width and average velocity from
the prior day’s aeration estimates to locate sampling sites.
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Once the stations have been located, take GPS readings for the injection location and the
sampling station and measure the distance between the two points. Note results on field data
sheet.
Sampling Site #2
(above downstream fence)
Sampling Site #1
(below upstream fence)
Upstream
Injection Site
(well-mixed)
Downstream Flow
Figure 4. Depiction of a SCALER experimental reach that outlines the location of the
injection point and the two pulse sampling stations within the reach.
During the experiment:
Mix the dry solute masses into the specified volume of stream water in the bucket or
garbage can. Note, it takes a substantial amount of time to get high concentrations of salt into
solution, it is generally better to start this the night before, at site if too heavy to carry. Wash
solute residue from the bags into the container with more streamwater. Using a container with a
completely open top may be useful to allow a truly “instantaneous” injection of the solute
solution. It may take some time for all of the material to dissolve, so mechanical shaking/stirring
of the solution can be used to speed this process. Set aside a small aliquot of the injection solute
for later analysis. This sample will need to be diluted prior to analysis, so a scintillation vial
(available volume for analysis) will suffice. It is important to keep this highly concentrated stock
solution separated from other samples to prevent contamination. Triplicate background samples
(dissolved nutrients and conductivity) should be collected at the injection station as well as the
two sampling stations prior to starting the experiment.
Install a conductivity (bromide or fluorescence) meter at each of the sampling stations in
order to monitor the arrival and departure of the added solute slug making sure they are in the
water column. Record the offset of the conductivity meters time and the recorders watch. If a
sonde or HOBO conductivity logger is available, log conductivity for later analysis/interpretation
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at short intervals (as short as 1 s, no more than 1 min). It is likely (especially for smaller streams
and probably at every top station) that the breakthrough curve for a slug addition happens too
quickly (sampling every 30s – 1 min) to allow for filtration in the field. In this case, it is better to
use a different set of field sampling bottles (e.g. wide-mouth, 250 mL Nalgene HDPE bottles).
Number these bottles sequentially simply record the bottle number in the field datasheet with the
time and conductivity (note time in 24-h format). It will be easiest if you fill the temporary field
sample bottles in numerical sequence from 1 to n. These temporary field bottles can be returned
to the lab to filter into final sample bottles (60 mL). The method described below assumes that
temporary field sample bottles will be used, but if there is sufficient time for filtering in the field
(i.e. one person pulls up stream water in syringe while the other person filters water into bottles)
it is preferable.
Pour the well-mixed, completely-dissolved solution into the stream, in one very quick
motion, at the pre-determined mixing site. Be sure, however, to not pour the solution so rapidly
that tracer is forced into interstitial spaces in sediments. As you pour attempt to put most of the
solution into the thalweg, but roughly apportion the rest into the less rapidly flowing regions (i.e.
mix across the stream with the bulk of the solution going into the thalweg). Quickly rinse the
release container with stream water and put that in. Note the release time. At the two sampling
sites, monitor the conductivity using the conductivity meter, noting the rising limb, peak and
falling limb of the breakthrough curve.
In all cases, samples
should be taken in a well
mixed area (thalweg) facing
upstream or standing out of the
channel. In fact, on the day of
nutrient measurements, as
little disturbance as possible to
the streambed is critical. In
general at least two people
should be assigned to each
station, with one taking
samples and the other
recording time, conductivity
and supplying sample bottles.
More samples over
time are better than replicate samples at any individual time, statistically. Samples need to be
taken most quickly when concentration is changing rapidly, so you will need to sample very
quickly when the peak reaches the sampling station. Use this real-time data to guide the
collection of grab samples to ensure characterization of the entire concentration-time curve for
the biologically-active tracer as well as the conservative tracer.
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Plan to collect at least 20-25 samples during the breakthrough curve. Background
sampling before the peak starts to hit should be sparse, but with 3 baseline samples taken. As the
conductivity begins to rise, the most intense sampling will occur. A good measure of travel time
requires getting a good fix on when the peak comes through, so pay careful attention to sample
through this period. Sampling should cease after characterization of the falling limb of the
breakthrough curve, when conductivity has returned to baseline. However, do not stop sampling
prematurely at the tail end of the experiment. Measuring the tail is the only way to get a good
estimate of discharge from a pulse release (http://pubs.usgs.gov/twri/twri3-a16/pdf/TWRI_3A16.pdf) Be patient and reserve at least 2-3 samples for the long tail of the breakthrough curve.
Essentially, the sample times should be long till the peak starts, very often as the peak comes off,
and become less and less frequent as the peak has passed.
Take samples by either dipping the temporary field sample bottle in the stream to “grab”
a sample or with a syringe. Remember, record sample number, date, time and conductivity
(bromide or fluorescence) for each sample taken.
Once the experiment is complete (or during the experiment if you have time or extra
hands), filter all of the water samples. Ideally, the field is best, or filter in a clean lab as soon as
possible after the uptake is complete. Remember cleaning products, cigarette smoke, and
mowing lawns can lead to atmospheric ammonium contamination. Also, leaving the caps off the
bottle can cause rapid volatilization.
Arrange all of temporary field sample bottles on a lab bench in two sequences: one
sequence for the rising limb of the breakthrough curve and one for the falling limb. Filter the
samples from the two limbs separately, always working from the lower concentrations to the
higher concentrations near the peak. Filter each sample from the temporary field bottle with a
60-mL syringe through a GFF filter into a 60 mL bottle. Pour sample into the syringe closed with
a stop cocks, leaving some air space for the plunger. Since samples along a time series will be
closer in concentration to the next sample, there is no need to rinse and change filters between
samples, but you should pour 5 mL extra into each sample and force that through the syringe
before collecting the filtrate. You can use this first 5 mL to rinse the sample container as well.
Do not use the same filter for a lower concentration sample or for a sample from another limb of
the breakthrough curve. If samples are not analyzed immediately, freeze them. If you freeze
samples, be sure that you leave enough headspace for the bottles to expand as they freeze.
Collect data on stream widths after the release is completed. Conduct a width-depth
survey for at least 15 transects along the experimental reach. Width measurements are used to
calculate nutrient uptake parameters.
During the experiment:
Collect three water samples for background conservative tracer concentration. Collect
triplicate samples of concentrated solute solution in case you want to re-analyze this.
When releasing the slug solution, gently but quickly release the solution under water
without creating splashing or bubbling. We suggest you practice on a blank carboy of stream
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water prior to the first real release. You are trying to minimize disruption of the stream bottom,
make the release as close to instantaneous as possible, and mix through the channel as much as
possible. Record the release time on the field data sheet. Samples must be collected at a precise
timing. At the sampling, begin sampling nutrients and inert solutes as soon as the conductivity
begins to increase.
The timing of samples varies depending on the time required for the break-through curve
to pass the sampling station. Strive to collect at least 20 – 25 samples over the break-through
curve. On the rising limb of the conductivity curve, samples will need to be collected more
frequently to capture the rapid change in tracer concentrations. Samples must be collected at a
precise timing using the timing intervals discussed previously. Always collect a big enough
sample for later quantitative analysis of both ammonium and inert solute, including extra for
duplicate analyses if the first round fails for some reason.
Continue sampling until the conductivity in the stream is within 2% of background level
3.7.
Water chemistry sampling of experimental reaches and synoptic sites
Material list
Sample bottles, labeled (~110 125 mL Nalgene acid washed, for one year and biome, note
minimum requirement for acid washing is 1 h soak in 0.1 molar HCl)
Syringe and filter apparatus
Filters (GFF from Zefon)
Forceps
Cooler with ice
Water samples are collected at the beginning and end of the experiment as well as days
11 and 22 of the experiment. Samples are collected below and above of the upstream and
downstream fences, respectively, of the control and treatment reaches. At each location, 125 mL
of filtered and unfiltered stream water each are collected and place in the cooler and frozen
immediately upon return to the laboratory. Additionally, water chemistry samples are collected
once in the middle of the reach at all 20 synoptic sites during the experimental period.
3.8.
Habitat delineation of experimental reaches and intensive synoptic sites
Material list
Field data sheets
Pencil
High-resolution global positioning system (GPS)
Digital camera
Gravelometer
Rangefinder or measuring tapes (50 or 100-meter)
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These data will need to be collected to inform modeling attempts to upscale to network
from reach-scale measurements. As this could be a substantial amount of work, this section
might be modified once it has been attempted at the first few sites.
Use the pre-synoptic sample collection as a guide to establish the substrate size
distribution. Refer to Table 4, the modified Wentworth scale, to delineate substrate size classes.
Walk along the synoptic reach (intensive sites only) or the experimental reaches and sketch the
distribution of the different size classes. Once the sketch is completed, begin delineation with the
high resolution GPS.
Create a new shapefile in ArcPad for the stream outline. Walk the stream starting at the
upstream end moving downstream on the right bank and back upstream on the opposite bank. In
a second shapefile delineate the riffle, run, and pool habitats which the habitat exclosures are
based on. In a third shapefile, delineate the substrate patches that were ID during the sketching.
Make sure GPS preferences are set to XX points and XXm.
3.9.
Whole-stream metabolism at synoptic sites
Material list
6 4ft ½ in rebar
Mallet
6 YSI ProODO meters with extended battery compartment
6 Odyssey irradiance meters
Bull’s eye level
Zip ties
Bucket (calibration)
Aerator (calibration)
Whole-stream metabolism at the six intensive synoptic sites (i.e., network scale) is
measured using a single station method. One DO and PAR meter are place in each site and
deployed for a minimum of 24 hours. Procedures and protocols follow those detailed at the reach
scale (3.4.). If possible, the six sites should be surveyed as close together in time as is possible,
but also feasible.
3.10. Nutrient uptake at synoptic sites
Nutrient uptake and SF6 reaeration measures at the intensive synoptic sites (n=6) follow
the methods detailed for experimental reach sites (section 3.5. ). The reach length for
measurement of nutrient uptake is determined with the initial SF6 solute release as for
experimental reaches (section 3.1). Reach length should be determined in part by selecting a
well-mixing release site, a constrained top station close to the release site, and a downstream
station at another constriction. Reach length should be determined similarly to the methods
detailed for the reach sites based on travel time (section 2.2).
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3.11. Algae chlorophyll a and benthic organic matter (BOM) sampling at synoptic
sites
Material list
Loeb sampler with brush (4.9 cm2 area) for bedrock and substrates with large planar area
Neoprene template (4.9 cm2 area) and toothbrush for smaller substrates
Small PVC cylinder (4.9 cm2 area) and a thin, stainless steel standard non-slotted spatula (the
kind you use for a frying pan, not the kind for spreading icing or cleaning out jars) for sampling
soft substrates
White tray
Whirlpacks with labels
Squirt bottle for rinsing
Cylindrical sampling templates (several pipe-corers see below)
Meter stick
Sample bags for leaves and wood (10 per reach, 1 per sample)
Sample bottles for FBOM subsamples, approx. 250 mL size (10 per reach, 1 per sample)
Small plastic ruler
Cooler
Strategy:
We want to account for as much of the primary producer biomass as possible and are up against
several constraints. Scraping misses 20-50% of attached chlorophyll depending upon the
roughness of the substrata sampled (Murdock , J. N. and W. K. Dodds. 2007. Linking benthic
algal biomass to stream substratum topography. Journal of Phycology 43:449-460). However,
large rocks are difficult or impossible to sample with whole rock extraction. At the same time
with fine substrata such as sand or silt, whole substrata extraction is necessary. Thus, we will
combine whole substrata extraction with scraping of large substrata. In sites where most rocks
can be extracted (e.g. there are few or no rocks), only whole-rock extraction will be used.
However, if people are interested in the proportion of easily removed chlorophyll that is
influenced by consumer manipulation, they may still want to scrape and extract whole rocks.
Calibration measurements will be made of intermediate sized substrata that are scraped and
whole-rock extracted both to correct for the inability of scraping to remove all chlorophyll. If
possible, whole-substrata extraction is the preferred method. While scraping may simulate what
herbivores are capable of, this project is centrally concerned with scaling metabolism as
influenced by consumers, and only sampling the portion of the primary producer community that
is susceptible to grazing could substantially over estimate the effects of grazers while seriously
underestimating the total producer biomass. If we overlap whole rock extraction and scraping on
some comparable substrata, we can then test the hypothesis that rougher rocks will be less
influenced by changes in the herbivore community. These comparisons can be done along with
chlorophyll sampling or as side trials at roughly the same time as chlorophyll sampling.
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As our initial site surveys have accounted for the amount of cover of major substrata in pools and
riffles, the strategy will be to expend the most effort sampling the most common substrata.
Selection of sampling areas
Each synoptic reach is divided into 10 blocks (i.e., 10 m per block if reach length is 100 m), with
blocks moved such that they coincide with the beginning of riffle or pool habitats. The
proportion of riffle and pool area over the entire reach are used to stratify the sampling effort.
Proportion Riffle
0 – 10%
10 – 30%
30 – 50%
50 – 70%
70 – 90%
90 – 100%
# of Riffle Areas Sampled
0
2
4
6
8
10
Use random numbers to decide the perpendicular stream transect location within a block (1-X m
for meter markings, and 1-10 for location across the transect). Within the randomly selected
transect, sample the dominant substrata types for both biofilm and BOM (n=1-3). Because the
area sampled for biofilm with the Loeb sampler is small, 3 sub-samples are taken at each transect
location and combined for a single sample in a Whirlpack bag. The substrata sampled are
determined by the upto three substrate types that make up over 80% cover. The type of habitat
sampled is recorded as is a GPS location for each sample.
Biofilm sampling:
All samples are stored in the dark in a cooler as soon as they are collected.
Hard substrates: On large rocks and bedrock, use the Loeb sampler (4.9 cm2 area). Press the
neoprene firmly against the substrate, then use the brush to vigorously scrape the substrate.
While maintaining contact with the substrate, pull the brush handle up, so that the water/biofilm
slurry enters the Loeb sampler tube. Cover the hole at the base of the sampler to maintain suction
and slide the spatula between the neoprene and the rock. With the other hand hold the Loeb
sampler to the spatula then lift Loeb sampler from the substrate above the water and invert. If
you have clearly lost more than 1/5th of the sample, discard and start again. From here empty the
contents into a Whirlpack using a squirt bottle to thoroughly rinse all biofilm into the collection
container. For each sampled ‘hard substrate’ area in the stream, use 3 Loeb and/or template
samplings for a composite (total area sampled 14.7 cm2), collecting material from all 3 scrapings
into the same bottle or Whirlpack bag.
If the hard substrate can be easily removed from the stream, place the substrate in the sampling
tray and scrape/brush an area the size of the template. Rinse and collect the material.
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For rocks that are about the same size as the Loeb area, simply remove three and place in a
whirlpack bag for later whole-rock extraction.
For smaller gravel substrata, place template on stream bottom and collect all pieces of gravel on
the surface to approximate the size of the template and place them in a whirlpack bag for later
extraction in ethanol. We will use ethanol as the hot ethanol method has been shown to offer
superior extraction to acetone and grinding methods (Sartrory and Grobbelaar 1984).
To allow correction of Loeb and scraped samples for missed chlorophyll we can compare rocks
that are similar to the dominant type of larger rock (with respect to smoothness). Pick 6
individual rocks around the size of the template. Place each rock in the pan and scrub with the
brush the same was the template and Loeb samples were scrubbed. Rinse and save the scrubate
and save each rock separately for chlorophyll.
Soft substrates wood and leaves:
Soft substrates are sampled with the PVC tube. Place the PVC tube on top of the biofilm/FPOM
associated with depositional habitats to a depth of 2 cm. Slip the spatula underneath the
substrate, get a seal with the PVC and lift and dump contents into the sample tray, transfer to a
Whirlpack bag. Take 3 samples for a given location, collecting material from all 3 collections
into the same Whirlpack bag. If wood or leaves dominate and need to be sampled, then take three
pieces roughly the size of the Loeb and template and place in a whirlpack bag.
Upon return to the laboratory, rocks, leaves, and gravel can be frozen. Fine sediments should be
settled if excess water is present, the water decanted and then they can be frozen. Liquid samples
from scrapes need to settle overnight in the refrigerator or be well mixed and filtered. The settled
material and/or filter are to be kept frozen until analysis.
Analysis for chlorophyll should be completed within one month if at all possible. Data show 510% loss of chlorophyll over 6 weeks of freezing Hallegraeff, G. and S. Jeffrey. 1985.
Description of new chlorophyll a alteration products in marine phytoplankton. Deep Sea
Research Part A. Oceanographic Research Papers 32:697-705.
.
BOM sampling:
We probably need several cylinders to sample this in most streams, depending upon the depth of
the stream that can be sampled. Given the method the longest cylinder should be no longer than
10 cm shorter than the reach of the longest arm in the group. Although long cylinders are
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necessary for sampling deeper pools, they are unwieldy so you will also want a shorter cylinder
(say 16”). Cylinders should have a diameter of at least 8” but probably not much wider than 12”.
You can use furnace duct material (line top of cylinder with split rubber tubing or you will cut
yourself) or PVC (bottom of cylinder should be sharpened either by a shop or file or dremmel
with a cutter to get sharper). The diameter of each cylinder should be measured.
For every one of the 10 areas that has fine enough sediments that it can be sampled, take one
sample. For each sample, place the cylinder as deeply into the sediments as possible. Remove all
coarse benthic organic matter (CBOM, > 1 mm, leaves and wood) within the cylinder and place
in a durable plastic bag (e.g., pollination bag). Collect fine benthic organic matter (FBOM, <
1mm) to a depth of 6 cm (to approximate baskets) by swirling water and 6 cm deep layer of to
fully suspend all sediments inside the cylinder. While swirling, quickly collect a sample of the
suspension in a bottle and place in the cooler. It is easiest if you use a graduated beaker and
collect the same sample volume every time. Measure the depth of water in the cylinder in order
to calculate the total volume of suspended surface FBOM (record depth at 4 locations within the
cylinder and then use the average). This volume is used together with the mass of the filtered
subsample to compute the FBOM mass per unit area (area within the cylinder). For these
samples you will need to record GPS location, habitat, diameter of cylinder, and 4 depths for
each sample taken.
3.12.
Metabolism and nutrient uptake of patch exclosures
Material list
7 chambers (bring one as reserve)
7 covers
Electrical boxes
12v battery
7 YSI ProODO meters (one as reserve)
Odyssey irradiance meter
Tools and spare parts (Allen wrenches, screw drivers, duck tape, assortment of screws etc
supplied with chamber)
1L volumetric cylinder, bucket with 8L mark
Funnel
Ammonium stock solution calculated so that 3 mL will take 10 L to 3 x background
concentration or at least 25 ug/L NH4-N
Sample bottles (85 at least 30 mL bottle size, acid wash nalgene)
Sample syringe and filter apparatus (GFF Zefon)
5 pieces black window screen large enough to cover a chamber
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Figure 4. Motor shaft parts and assembly drawing for chambers.
Motor box assembly:
Before beginning assembly of motor housing, be sure to review the video posted to dropbox
(“Fixing motor in motor assembly”, Manual folder) and check that all parts are available from
shaft assembly diagram (Fig. 4). In order to simplify the assembly process, certain pieces have
been pre-assembled. This includes: the propeller will be connected to the propeller connecter, the
PVC tubing sleeve and rubber stopper will be on the brass sleeve, and the motor connecter will
be attached to one side of the brass drive shaft.
Begin assembly by attaching the post to the top of the motor box. The bottom bolt, metal washer
and plastic washer should be removed from the post, leaving one metal washer and bolt attached.
Place the post through the smaller of the two holes on the top of the motor housing box. Next reattach the plastic and metal washers and bolt. Tighten the bottom bolt with a 7/16 wrench until
snug (over tightening may crack plastic), making sure that the loop at the top of the post is
parallel with the longer side of the chamber. Next put the brass sleeve (H) with attached rubber
stopper (L) into larger hole on the motor box. It is easiest to attach stoppers by simultaneously
twisting and pushing down. Make sure that the pink PVC tubing (G) is in line with the white set
screws in the plastic shaft. Adjust accordingly by moving brass sleeve up or down within the
rubber stopper. Next remove the motor connecter (J) from the brass drive shaft (F) and slip brass
drive shaft into brass sleeve. Place the motor box on its back, and attach propeller connecter (D)
to brass drive shaft by tightening the set screw (E). Be sure to leave enough space on the top of
the shaft for the nylon spacer and the motor connecter to attach. Also check that the set screw on
the propeller connecter will attach to the flattened area of the brass shaft. While the motor box is
still on its back, slip on the nylon spacer (I) and attach the motor connecter to the other side of
the brass shaft. One of the set screws on the motor connector should be tightened to keep the
brass shaft from taking up too much space within the connecter (i.e. this will ensure there is
enough space to attach the motor). Again be sure that the set screw will be attached to the flat
area of the brass shaft.
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Put the motor box upright. Next tighten (or loosen) white set screws until the shaft is held in
place, while the propeller is also spinning freely within the PVC elbow. The top of the propeller
should be flush with the top of the PVC elbow (this might vary, but most of the propeller should
be contained within the elbow). At this point, it is vital that the propeller does not make
contact with the PVC elbow when spinning the brass shaft. Attach motor (K) to motor
connector by loosening the top set screw, inserting the flattened edge of the motor shaft, and retightening the set screw. Finally loop the rubber band (attached through the loop of the post) over
the motor. The rubber band should loosely hold up the motor; keeping the motor in place, while
still allowing some wiggle room during operation.
Filling and starting the chamber:
Before assembly of chambers, be sure to review the video posted to dropbox (“Filling
chamber…”, Manual folder). Fill inner box with water and stopper the hole. Place on stilts inside
chamber with the stopper down and towards the back corner of the chamber. It will need to slide
under the two tabs on the sides of the chamber. Next slide the table that the baskets will sit on
under the lip of the box making certain it fits flat on the stilts and is held in by the two stops on
the motor side. Place entire motor housing assembly into chamber and tighten link locks.
Ascertain that the chamber drain stopper is in place. Lay out the brown cover and place chamber
on top with the motor assembly slightly elevated relative to the other end of the chamber. The
portal for the stopper should be at the highest point of the chamber. Fill a bucket to the 8 L mark
and slowly pour the water into the chamber. Test that the propeller is still moving freely.
When starting the motor, make certain the power supply is to the off position. Attach the power
supply to a 12v battery. BE VERY CAREFUL TO ATTACH RED TO THE POSITIVE
TERMINAL OR YOU WILL INSTANTLY FRY THE POWER SUPPLY. Plug the wires
into the power box then clip onto the motor (red to plus). Slowly turn up the power as not to strip
the shaft connectors. The water movement should be down through the elbow to the far side of
the chamber and flowing back across the top toward the motor. If the propeller is running
backwards switch the connections from the power supply to the motor (but never at the battery).
The motor should be running quietly at this point. If motor is noisy, adjust the rubber band and if
this does not help make certain propeller is still spinning freely.
Once the motor is running, set velocity. To set velocity take a multimeter with leads plugged in
to measure amps. Clip the ground lead to the positive red lead from the power source, and the
red text lead to the + poll on the motor. Turn the meter to read up to 2 amps. Turn up the motor
till the multimeter reads 0.5 amps. This will give us an approximate velocity in all chambers of 7
cm/s. This value also lets us know what batteries we need and how often. 12V lawn mower and
scooter batteries have 18 amp hours. Modest size deep cycle marine batteries have 40 amp hours.
If we run 6 chambers for 8 hours we would have 24 amp hours, so 2 lawn mower batteries
charged per day should run the chambers. Also note as batteries drain their voltage drops, so
every couple of hours we need to reset the velocities by checking the amps, because the
amperage will drop as voltage drops.
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Gently as possible collect the three baskets from the exclosure using the chamber lid as a
transporting device (the other two baskets will be used for FBOM, chlorophyll and
invertebrates). Gently lower the sediment baskets to be measured onto the table between the box
and the motor assembly. Then attach the lid to the chamber, starting with the link locks
connecting the motor assembly to the lid. Then close the link locks moving along the length of
the chamber toward the end opposite the motor housing. Continue to add water through the
portal for the oxygen probe using a volumetric cylinder and noting the volume until the chamber
is nearly full. Insert the oxygen probe and twist in place. Make certain temperature sensor for
oxygen probe is shaded inside the chamber or it will give a bad measure. Using a funnel fill in
the rest of the chamber through the stopper portal and close with stopper when full. If the flow
dislodges more bubbles they should accumulate at the stopper portal and more water can be
added to try and get rid of most bubbles, a few small ones will not matter. While the chambers
are shipped leak-free, small leaks do not matter much as a 100 mL loss over the 10 minutes or so
of measurement would be only a 1% volume loss, and a 10 mL per minute leak is a bad leak.
Pre-sampling metabolism measurement (one day in advance): Try to do this measurement in full
light midday. Use spare baskets (which have been incubated for the experimental time, or
alternatively have been filled with similar substrate from the treatment reach) and collect some
substrate from the stream for this method test. Continuous measurement of PAR (using an
Odyssey irradiance meter) is measured to allow for normalization of differences due to diurnal
metabolic characteristics. Close the cover over the chamber and the light logger and start logging
dissolved oxygen at 1 minute intervals. Record starting time and DO concentration and log until
dissolved oxygen has changed by 1 mg L-1 from the initial concentration. Alternatively, plot on
the fly and determine if you get a straight line decrease. Stop the DO meter, remove the cover,
and restart the DO meter. Record for the same amount of time required to get a significant
change DO drop in the dark (if NEP = then actual DO might not change at all in this light
incubation). Plot data for ER in using the dark measures and NEP using the light measures to
determine how long of a measurement is required to get a consistent change in O2 over time. The
R2 should be larger than 0.95. The point is to use the smallest amount of time per measurement
to get a statistically significant result. The absolute minimum time is how long it takes the O2 to
completely mix in the chamber (less than a minute) but at least five points and thus minutes are
needed to get a regression. The longer the time used to take the measurements the more the
stream water in the chamber deviates in temperature and nutrient content from the stream. The
determination is made using ER because if NEP is close to zero in the light, there will be no
detectable change in O2 concentration.
Plot changes in dissolved oxygen in both the dark (ER) and the light (NEP) to test for
linearity. If changes in dissolved oxygen are linear. We will use at least 10 minutes even if we
can get a good relationship over less time.
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Once the time needed to measure a disenable change in DO (e.g., 10 min for linear) is
determined, the photosynthesis irradiance curve needs to be determined. This curve is required to
account for changing light levels when the full set of baskets is measured throughout the next
day. Make certain you are logging light while you are making the measurement. Cover both the
chamber and the logger with 5 layers of screen and log O2 for the amount of time determined in
the respiration. Remove 1 layer and repeat the procedure. Continue repeating up to full light (you
will generate rates for dark (initial), 5, 4, 3, 2, 1, and 0 layers of screen). Be logging PAR ding
this time so you know what light there is under each of these light conditions.
Uptake measurements: For nutrient uptake measurements, add 3 ml of the nutrient stock solution
after the dark/light incubations for metabolism are completed. Ammonium is added to increase
concentrations to 3x background. This is close to background but high enough to allow analyses.
If your concentrations are so low that you are not confident that this will work, shoot for about
25 µg NH4-N/L. Take the first sample after 1 minute and then space the rest of four samples
over the course of 40 minutes. The removed water is not replace and some aerated headspace
will develop. If possible quickly analyze some runs to be certain there is a significant decline in
concentration. Water samples (30mL) are removed, filtered immediately into acid washed
bottles, and placed in a cooler. Water volume is replaced with air. Samples are frozen upon
return to the laboratory.
Experimental measurements: Carefully remove baskets from the exclosures, place on chamber
lid to transport, and follow instructions above. Site names are created based on naming
convention (see section 6.1.). Chl and BOM will come from extra baskets.
Emptying chamber: Turn power supply off and unplug from battery. Open lid and let some of the
water run out over top. Gently remove baskets, and empty them out. Pull the drain plug since
lifting a full chamber runs the risk of dropping and cracking the whole thing. Remove motor
assembly and rinse chamber with fresh streamwater. Check tightness of all set screws and
replace motor assembly, and the chamber is ready for the next filling and sampling. Check
amperage if it has been a couple runs without it.
3.13. Algae chlorophyll a and benthic organic matter sampling of patch exclosures
and experimental reaches
Material list
Whirl packs (50), labeled
Cooler with ice
Nalgene bottles (50 500 ml), labeled
Bucket with 4 L mark
100% Formalin
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From one strawberry basket in each habitat exclosure as well as from baskets placed in the
pools and riffles of the control and treatment reaches, whole substrates are collected and placed
in whirlpacks. Make sure that the surface area of the substrate pieces collected cover at least 5
cm2 of the basket. Remove up to three rocks depending on substrate size from each basket. The
selected rocks are returned to laboratory in a cooler and frozen as soon as possible. Analysis for
chlorophyll should be completed within one month.
Fill the bucket to the 4 L mark with stream water. Place the remainder of the strawberry
basket in the bucket. Agitate to suspend benthic organic matter and take a 500 ml sample.
Samples are transported back to the lab and refrigerated if not processed immediately. If they
can’t be processed within two days, add 40 mL of 100% formalin (37% saturated formaldehyde)
to create a 8% formalin solution. Return to the laboratory for filtering. Processing should be done
within a week. Note, if you have time to process BOM materials that evening, formalin is not
necessary for this step.
It is important to don appropriate PPE while handling it (even the field… safety glasses and gloves at
a minimum). A good trick to do is to pre measure your 37% saturated formalin in to a scintillation vial
(e.g., 40mL) and have the concentrated formalin double-bagged to reduce risk of any environmental
release. Then, just dump the opened vial and cap into your container.
In addition, control and treatment reaches are sampled for chlorophyll a and BOM as
described for the synoptic sites (see section 3.9). Reaches will be divided into six squares, and
the stream length of a square will be noted. In each square the dominant substrates are taken
from preliminary sampling data and sampled. If habitats with fine sediments exist that are not
represented in the sampling, additional BOM samples are collected from those areas using the
core sampler (n=6). These samples should be taken to cross calibrate baskets with sampling
methods used in synoptic sampling.
3.14. Invertebrate sampling of patch exclosures and experimental reaches
Material list
White pan
250µm sieve (2)
Water squirt bottle
Forceps
Whirl packs (32) labeled outside and in
Bottle with 100% formalin (diluted with stream water)
One strawberry basket from each habitat exclosure is designated for invertebrate
sampling (n=16). Additionally, strawberry baskets filled with representative substrata were
deployed by burying them flush with ambient substrata in the stream bottom of pools and riffles
of the control (n=8) and treatment reach (n=8).
Samples are collected at the end of the experiment by carefully removing them from the
stream bottom while holding a 250 µm mesh bag or sieve just downstream of the basket and then
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around the basket as it is lifted through the water column. Baskets are then placed in a white pan,
along with contents of the mesh bag/sieve. To retrieve the contents of the mesh bag, any material
in the bag can be rinsed to the bottom using stream water. When material is condensed in the
bottom of the mesh bag, the bag can be inverted into the pan or over a 250 µm sieve and rinsed
clean with a squirt bottle filled with stream water. If rinsed over a sieve, the contents of the sieve
should then be rinsed in to the plastic bag. All contents of the pan should be rinsed thoroughly
and scrubbed over a 250 µm sieve. Contents of the sieve should then be condensed in one area of
the sieve and rinsed with a squirt bottle into a whirlpack. The final sample should have only
enough fluid to keep the contents submerged, and enough 100% formalin should be added to
make the fluid 8% formalin. Each sample should have two labels: one within the whirl-pack and
one adhered to the whirl-pack. The internal label should be a plasticized paper label (e.g., write
in the rain paper). The same information should be written in permanent ink on the outside of the
whirl-pack.
4. Laboratory procedures
4.4.
Reaeration SF6 samples
The vacutainers need to be processed as soon as returned to the laboratory. Measure the mass of
all vacutainers to calculate the volume of water sampled. The mass of vacutainers is consistent
among vials allowing for volume to be calculated from the difference in mass between sample
vials and empty vials. To determine the mass of empty vials, weigh five empty vials and
calculate the mean. Samples should be processed quickly, within a week if possible. However,
as a week is likely not feasible, a control vial with a known volume of SF6 should be prepared to
estimate loss through septa.
Measure SF6 from the headspace gas on a Gas Chromatograph with Electron Capture
Detector. Note that the concentration of SF6 in vials is high and will swamp the detector.
Samples will need to be diluted before injecting. This dilution is accomplished using a sealed
flask and injecting with a high accuracy syringe (e.g. Hamilton glass syringe). Care should be
taken to avoid carry over as high concentrations of SF6 tend to carry over.
After measuring SF6, the conservative tracer concentration can be measured on the water.
Discard used vacutainers as SF6 binds to the septa of vials and contaminates subsequent analyses
if vials are reused.
Laboratories at LUQ (University of New Hampshire, McDowell), KNZ (Kansas State
University, Dodds), and CPC (University of Alaska Fairbanks, Jones) can process samples and
use their standard operating procedures. Since relative values are key for analyses direct
comparison among laboratories is not required. If you cannot analyze your own samples, you
need to negotiate with another laboratory to analyze them for you. Generally they approach
should be to offer to analyze some of their samples in return (e.g. do BOM samples for them).
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4.5.
Streamwater nutrient and ammonium uptake samples
Streamwater samples are thawed and analyzed for soluble reactive phosphorous, ammonium,
nitrate, total dissolved phosphorous, total dissolved nitrogen, and dissolved organic carbon as
well as particulate nitrogen, phosphorous and carbon. Samples for total N and P can be
determined via whole-sample digestion of filtered samples and addition of values from filters
that are analyzed for N and P content, however keep in mind that this still requires a whole
sample digestion. The benefit is that particulate and dissolved N and P can be indicated. Samples
for whole-stream and chamber nutrient uptake are analyzed for ammonium only. All samples for
pulsed release uptake experiments should be analyzed.
Each biome and laboratory analyzes samples based on their standard operative procedures. To
compare values across biomes, all laboratories participate in the USGS round robin
(http://bqs.usgs.gov/srs/) to verify accuracy. The check samples are available two times per year,
and checks should be run at least once per year, though twice would be better, and running them
before you run your unknown samples is preferable so problems can be fixed before you analyze
samples.
4.6. Benthic organic matter (BOM)
Materials needed
Pre-ashed, pre-weighed GFF filters 45 mm
Pre-weighed aluminum tins
Filter apparatus
Graduated cylinder
Forceps
Paper bags
Scribe numbers into weighing aluminum tins and pre ash. Pre weigh enough tins to contain
CBOM samples. Tins and filters are pre-ashed at 450°C for a minimum of 3 hours. To pre-weigh
filters (need number) put numbered tin on scale, tare, and note weight and tin number on lab
sheet. Never handle ashed tins or filters by hand, use clean forceps.
Basket samples:
Coarse BOM (CBOM; > 1 mm): A volume of 500 mL of the benthic organic matter samples is
filtered through a 1 mm mesh. The mesh is then rinsed into a pre-weighed aluminum tin
Fine BOM (FBOM; 0.45µm – 1.0 mm): Use the filtrate from the CBOM sample and filter a
known volume through a pre-ashed and pre-weighed GFF filter, until the filter clogs. Record the
volume of filtrate. The filter is then transferred into a pre-weighed aluminum tin (fine; FBOM).
All tins are dried at 60° C for a minimum of 24 h, and then weighed. Tins are subsequently ashed
at 450°C for a minimum of 3 hours, and re-weighed. Dry mass, ashed mass, and ash-free dry
mass (AFDM) are calculated based on measurements, and converted to per area basis (10 cm2
surface area of basket). Save and dry an additional set of samples on filters if PP PC and PN are
desired.
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Corer samples: CBOM: Separate wood from leaves, place in small paper bags, and dry both at
60°C for a minimum of 48 h. Record dry weights of each group, then weigh and combust a
subsample of each for a minimum of 3 hours at 450°C. Save the remainder the use for nutrient
ratio analysis (see section 4.5.1.).
FBOM: Shake sample bottle well, and pour a subsample into a graduated cylinder. Record the
volume in the cylinder, and filter through a pre-combusted, pre-weighed GFF filter. The
remainder of the sample is used for nutrient ratio analysis (see section 4.5.1.). Dry filters at 60°C,
weigh the filters, and combust at 450°C for a minimum of 3 hours before re-weighing to
calculate ash-free dry mass and ashed mass.. To calculate FBOM standing stocks for each field
sample, multiply AFDM/volume filtered values by the total volume within the cylinder used in
the field, then divide by the surface area of the cylinder.
To determine standing stocks of each BOM compartment for the entire reach, weigh the mean
standing stocks for each habitat type by the relative proportions of each habitat type in the reach.
4.7. Chlorophyll a
Material needed.
GFF filters
131821010 Clavies® Autoclavable Bags 10X10 Bel art
50mL test tubes labeled with scratched numbers
95% Ethanol (probably lab will need to buy 5 gal drum)
Chlorophyll a will be determined by hot ethanol extraction using the method of Sartory and
Grobbelaar (Sartory and Grobbelaar 1984). Note hot ethanol extraction is far more efficient than
acetone. It the same as methanol, but methanol is more toxic.
Keep samples in the dark at all times, at least in low light when working on them. If the sample
is kept in the dark, only 1.3% of the chlorophyll degrades with the 5 min. hot extraction and 24hour storage. Use test tubes that have the tube numbers scribed on the side with a diamond pencil
(ethanol eventually degrades tube numbers written in sharpie, diamond pencils can be found at
Carolina Biological Supplies) for filter extraction.
Loeb and scrubbed samples: Thoroughly mix the biofilm-water slurry, then pipette and filter 2040 mL of subsample through a GFF filter (does not need to be pre-combusted or weighed).
Record filtered volume. Wrap each filter in a small square of aluminum foil, label, and freeze at 80°C until analysis. The remaining sample is used for nutrient ratio analysis (see section 4.5.2.).
All samples: Place the filter in a test tube (50 mL centrifuge tubes work well), fine sediments in
containers, and whole rocks, pieces of wood or leaves in autoclave bags. Add a known volume of
95% ethanol. Mark the location of the meniscus on the side of the tube, container or bag with
sharpie. If using tube or container, place a loose cap on top of the tube, if using bags fold over
the top, do not seal either. Heat the tube, container or bag in 79°C water bath for 5 minutes, then
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shake and keep cool (5-15°C) for 24 h in the dark. Tubes, containers and bags should be sealed
once cooled. Be careful, if ethanol spills on the side it can wipe out sample markings.
After extraction, use additional 95% ethanol to bring up to mark on side of tube if ethanol has
evaporated, then shake. Clear sample by centrifugation, filtration, or settling. Rocks in autoclave
bags can be shaken and allowed to settle for an hour before analysis.
There are three methods of analysis possible. As all methods are calibrated against the
spectrophotometric method, they all should work equally well. However, since most sites have a
spectrophotometer, this is probably the preferred method. There are three methods of analysis
possible. 1) Spectrophotometric with acidification correction for phaeophytin, 2) fluorometric
with acidification correction for phaeophytin or 3) fluorometric with specific lamp and filter
combinations to avoid phaeophytin. Options 2 and 3 require calibration by a sample analyzed
with method 1.
1) Spectrophototric method: Analyze sample with spectrophotometric analysis at 665 and
750 nm using a 1 cm spectrophotometer cuvette (APHA 2005). If adsorption is over 1.5
absorbance units, dilute sample. Add 0.1 mL of 0.1 N HCl for each 10 mL of extractant
after the first reading and let sample sit for 90 s to phaeophytinize all chlorophyll a before
reading. Amount of acid is important, too much causes precipitates. APHA (2005)
method is modified with absorption coefficients from Sartory and Grobbbelaar (1984)
Calculations are made as follows:
Chlorophyll a (mg m-2) = (28.78(6650-665a)*v/(A*l)
Phaeophytin (mg m-2) = 28.78 [1.72(665a)-6650]*v/(A*l)
Where, 6650 = absorption at 665 before acid addition with absorption at 750 nm
subtracted out, 665a = absorption at 665 nm after acid addition with absorption at
750 nm subtracted out, v = volume of extractant used (L), A = area of benthos
sampled (m2) and l = path length of cell (cm) (usually 1 cm).
2) Fluorometric method: Use a fluorometer with filters and lamps appropriate for
chlorophyll a analysis. Fluorometer needs to be pre-calibrated with chlorophyll solutions
of known concentration. Use 95 % ethanol to extract spinach leaves, a large amount of
filamentous algae, or some other chlorophyll source. Dilute samples and use method
above (the spectrophotometric method) to measure and calculate concentration of diluted
samples. Then place the diluted samples in the florometer, read fluorescence units, add
Add 0.1 mL of 0.1 N HCl for each 10 mL of extractant after the first reading and let
sample sit for 90 s. The difference in the values should be regressed against the calculated
chlorophyll concentration of each solution to create a calibration curve for the
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flourometer. Then unknown samples can be acidified and read. Flourometers generally
hold calibration for months, but the calibration samples will degrade rapidly, so to check
calibration you need to re-prepare fresh standards.
If you have the appropriate filters you can measure chlorophyll on a fluorometer
without the acidification step (Welschmeyer 1995, APHA 2005). However, you still need
a set of known diluted chlorophyll standards to calibrate the machine.
Determination of area sampled: If Loeb sampler was used, or known area was scrubbed, surface
is known. For whole rocks, directly scan the rock or trace the rock onto paper with pencil and a
known length or area and scan the area of the rock later. Use commercially available or free
image analysis software (e.g., ImageJ) to determine area. Some plant biologists have leaf area
measuring equipment, which is essentially a scanner and computer program to determine area
automatically. To get chlorophyll a per unit area of stream bottom (we do not try to account for
every nook and cranny of rock surface), calculate the total amount of chlorophyll a in the
extractant based on its volume considering any dilution, extrapolate absolute amount if
subsampling occurred and divide by area sampled as determined above.
4.8.
Nutrient ratio analysis (C:N:P)
4.8.1.
Benthic organic matter C:N:P
Five samples of BOM are analyzed, one from every other block (see section 3.10.). For CBOM,
collect a representative subsample, approximately 2 g each of oven-dried leaves and wood.
Wearing latex gloves, crush leaf litter with your hands inside the paper bag, then place
subsample of crushed leaves into a ball mill (a coffee grinder can alternatively be used) and grind
material into a fine powder (consistency of flour; approx. 3-5 min). Wood subsamples should be
cut up into small wood fragments using PVC pipe cutters. Place wood fragments into a ball mill
or coffee grinder and grind material into a fine powder (approx. 5-10 min). For FBOM, place
remaining volume into a labeled plastic scintillation vial, freeze at -20°C, freeze dry, and store at
room temperature until C:N:P analysis.
4.8.2.
Biofilm C:N:P
Five biofilm samples from each synoptic site are analyzed for C, N, and P content, one from
every other block (10, 8, 6 etc.) throughout the stream reach. For these, place remaining volume
of slurry into a labeled plastic scintillation vial, freeze at -20°C, freeze dry, and store at room
temperature until C:N:P analysis.
4.8.3.
Analytical analysis
Carbon, N and P content are determined using standard operating procedures from each
laboratory.
4.9.
Invertebrate processing
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5. Timeline of measurements
5.4.
Project timeline of field measurements (among years and across biomes)
Biomes will be sampled consecutively (Fig. 4, Table 6). Sampling will start at the
Luquillo (PR) LTER in January/February (proposed state date Jan-28), followed by Coweeta
Hydrological Station in March/April (proposed start date Mar-15) and the Konza LTER in
May/June (proposed start date May-5), and the two Alaska biomes with Caribou/Poker Creeks
LTER in June-August (proposed start date Jun-24) and the Arctic LTER in July/August
(proposed start date Jul-6). Caribou/Poker Creek (CPC) and the Arctic (ARC) are sampled with
overlap, meaning the setup and initial sampling is conducted in CPC, followed by set up and
initial sampling at the ARC site, before returning to and finishing measurements in CPC
followed by ARC.
Figure 4. Proposed timeline across biomes.
Table 6. Field postdoc travel schedule 2013.
Task
Begin End
LUQ site selection, prep
1/15/13 1/27/13
SCALER LUQ
1/28/13 3/9/13
LUQ to CWT travel, CWT site confirmation 3/10/13 3/14/13
SCALER CWT
3/15/13 4/24/13
CWT to KNZ travel, KNZ site selection
4/25/13 5/3/13
SCALER KNZ
5/4/13 6/13/13
KNZ to CPC travel, CPC site selection
6/16/13 6/23/13
SCALER CPC
6/24/13 8/3/13
SCALER ARC
7/6/13 8/15/13
In each biome, three experimental sites are sampled in 2013 and a new set of three sites
in 2014. The 20 synoptic sites remain the same in 2013 and 2014 to account for inter-annual
variation. Six synoptic sites are designated as intensive, meaning that whole-stream metabolism
and nutrient uptake will be measure at those sites. Additionally, the first three intensive synoptic
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sites are surveyed for consumers in 2013 while the next three sites are surveyed in 2014. The
experimental duration is 30 days, for a total of 41 field days to account for installation time
across experimental reach sites. All synoptic sampling occurs during the experimental window,
as close together in time as possible except for pre-synoptic sampling.
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5.5.
Experimental timeline (1 year and biome)
The timeline is detailed in the table below. Different colors represent the three experimental sites (labeled E1, E2, and E3 for
simplicity) sampled in one year at each biome. Each site requires a three day window for installation and initial measurements. All
sites also have a three day window for final measurements.
Pre-experimental preparation include the following:
1) Selection of sites: three experimental sites and 20 synoptic sites (with 6 designated as intensive)
2) Marking locations (2.2.) for fences (2.2.) and habitat patches (2.1.)
3) Build wood frames for habitat exclosures (2.4.1.)
4) Prepare materials for exclosure and sampling (including sampling bottles, whirlpacks, nutrients, fencing, rebar, …)
Day 1
E1 – Fence installation
(2.4.2.), consumer survey
(treatment reach; 3.1.)
Day 2
E1 – Consumer survey
(treatment and control
reach; 3.1.), patch
exclosures (2.4.1., 3.2.)
Day 6
Day 7
E2 – Water chemistry (3.6.), E3 – Fence installation
WS nutrient uptake (3.5.), (2.4.2.), consumer survey
start DO and light (3.4.)
(treatment reach; 3.1.)
Day 11
Day 12
E2 – collect DO and light
probes (3.4.)
Day 3
Day 4
E1 – Water chemistry (3.6.), E2 – Fence installation
WS nutrient uptake (3.5.), (2.4.2.), consumer survey
start DO and light (3.4.)
(treatment reach; 3.1.)
Day 5
E2 – Consumer survey
(treatment and control
reach; 3.1.), patch
exclosures (2.4.1., 3.2.)
Day 9
Day 10
E3 – Water chemistry (3.6.), E1 – collect DO and light
WS nutrient uptake (3.5.), probes (3.4.)
start DO and light (3.4.)
Day 8
E3 – Consumer survey
(treatment and control
reach; 3.1.), patch
exclosures (2.4.1., 3.2.)
Day 13
Day 14
E1 – Water chemistry (3.6.),
consumer survey treatment
reach disturb treatment
reach (day 11; 3.1.)
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Day 15
E3 – collect DO and light
probes (3.4.)
SCALER methods Jan 9 2013 v 1.0
Day 16
Day 17
E2 – Water chemistry (3.6.),
consumer survey treatment
reach (day 11; 3.1.)
Day 21
Day 22
Day 18
Day 19
Day 20
E3 – Water chemistry (3.6.),
consumer survey treatment
reach (day 11; 3.1.)
Day 23
Day 24
Day 25
E1 – Water chemistry (3.6.),
consumer survey treatment
reach (day 22; 3.1.)
Day 26
Day 27
Day 28
Day 29
Day 30
E1 – start DO and light
E2 – Water chemistry (3.6.),
E2 – start DO and light
E3 – Water chemistry (3.6.),
(3.4.)
consumer survey treatment
(3.4.)
consumer survey treatment
reach (day 22; 3.1.)
reach (day 22; 3.1.)
Day 31
Day 32
Day 33
Day 34
Day 35
E3 – start DO and light
E1 – Water chemistry (3.6.), E1 – Baskets: DO, nutrient E1 – Consumer survey all
(3.4.)
WS nutrient uptake (3.5.), uptake, chla, BOM, inverts reaches (3.1.)
end DO and light (3.4.)
(3.3., 3.8., 3.9., 3.10.)
Day 36
Day 37
Day 38
Day 39
Day 40
E2 – Water chemistry (3.6.), E1 – Baskets: DO, nutrient E2 – Consumer survey all E3 – Water chemistry (3.6.), E3 – Baskets: DO, nutrient
WS nutrient uptake (3.5.), uptake, chla, BOM, inverts reaches (3.1.)
WS nutrient uptake (3.5.), uptake, chla, BOM, inverts
end DO and light (3.4.)
(3.3., 3.8., 3.9., 3.10.)
end DO and light (3.4.)
(3.3., 3.8., 3.9., 3.10.)
Day 41
• Solid background colors indicate different experimental sites, 4 person crew (3 only day 13-30)
E3 – Consumer survey all
• Gray background designated for synoptic sampling
reaches (3.1.)
• Checking of exclosure fence should be conducted as often as needed and possible.
• Post-experiment clean-up: remove all exclosures and fencing
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6.
Data procedures – data manager
6.4. Labeling protocol
6.4.1. Field labeling
Labeling files are available for all samples that need to be collected and stored. Each file
represents ONE reach-scale experimental OR synoptic site. Biome and site names in
conjunction with date are unique identifiers. The naming convention is as follows:
1) Three letter code for biomes: Luquillo (LUQ), Coweeta (CWT), Konza (KNZ), CaribouPoker Creeks (CPC), and arctic (ARC). These will be used at the top of a label with
SCALER to detail the project. In labeling files, use the replace function to change ZZZ to
the appropriate letter code.
2) One letter code for experimental (E) or synoptic (S) sites
3) Number for experimental (1-6) or synoptic (1-20) identifier, the designation of which is
up to each biome, based on their preference (e.g., up-to-downstream, based on timeline).
Suggest to use E1 – E3 for 2013 however. In the labeling files, please replace the letter J
(should not be confused with anything else) with the appropriate site number.
4) In case of experimental sites, one letter code for reach type: Control (C), Patch (P),
Treatment (T).
5) For reach scale measures, top for blow the upstream fence, and bottom for above the
downstream fence, if applicable.
6) For baskets in the reaches, the designation is Location, and numbers 1-8 assigned
downstream to upstream.
7) For patch scale measures, use eXclosure (X) to indicate the exclosure and the numbers 1
to 8 to indicate which exclosure it is, labeled downstream (1) to upstream (8). The habitat
designation of each exclosure is noted separately of these lables.
8) For eXclosures, enclosed sides are called In and open sides are called Out.)
9) Date uses three letter codes to avoid confusion. Please replace 99 with the day, and Yyy
is month [please use letter code as in the first three letters of a month]).
For Example:
SCALER_KNZ
E2_C_top_X1_IN
Electronic sample labels are available for:
Water chemistry (waterchem_2013.docx)
Ammonium uptake (uptake_2013.docx)
SF6 (reaeration) (uptake_2013.docx)
Chla (baskets_2013.docx)
BOM (baskets_2013.docx)
Either use Waterproof, sticky labels work well and can be purchased at: http://www.staples.com/Avery5520-White-WeatherProof-Address-Labels-1-inch-X-2-5-8/product_440728?externalize=certona
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Or paper labels put on with clear packing tape.
6.5.
Field data sheets
Data collected in the field is noted in notebooks. Such data include notes on weather and stream
conditions, goals for the day, people also present in the field. In terms of data collection they may include
placement of loggers and meters, macro-consumer surveys (species, lengths), and stream widths
depending on what the method sections call for. All field notebooks and field sheets should be write-inthe-rain type paper.
6.6.
Electronic data sheets
6.6.1.
File naming protocols
The table below lists all the file names and a description of their content as well as the suggested
interval for submitting (e.g., timing). The first part of each file name is the biome abbreviation
followed by the current date in the format of YYMMDD. The second part of the file name
indicates the metric contained in the file followed by the scale at which the data were collected.
All files are available in the templates folder under Data in the SCALER all dropbox. Please use
these templates since they are the basis for the database incorporation of the data.
Table X. File names of raw SCALER data.
File name
Content
XXX999999_habitat_synoptic.csv
Location codes, date,
physical parameters
XXX999999_habitat_reach.csv
XXX999999_sitedescription_all.csv
XXX999999_widths_synoptic.csv
XXX999999_widths_reach.csv
XXX999999_light_synoptic.csv
XXX999999_light_reach.csv
Specific mapping
information of
experimental reaches
GPS information and site
description
Location codes, date,
widths
Timing
Potentially twice per biome
and year, from initial
survey before experiment,
and during experiment
Once per biome and site
(can be combined in one
file)
Once per biome and year
Potentially twice per biome
and year, from initial
survey before experiment,
and during experiment
Location code, date, widths Once per ammonium
uptake, likely combined to
one file at the beginning
and end of the experiment
10-min interval data from
Once at 6 synoptic sites
light loggers, min. 24 hours during experiment (sites
can be combined in one
file)
10-min interval data from
Beginning and end at each
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SCALER methods Jan 9 2013 v 1.0
ProODO, up to 5 days
continuous
XXX999999_light_patch.csv
XXX999999_DO_synoptic.csv
XXX999999_DO_reach.csv
XXX999999_DO_patch.csv
XXX999999_DO_synoptic_cal.csv
XXX999999_DO_reach_cal.csv
XXX999999_DO_patch_cal.csv
XXX999999_DO_lineritylight.csv
experimental site
experiment (sites can be
combined in one file,
suggest one beginning one
end file)
1-min interval data from
One file per experimental
Odyssey light logger,
reach site, can be
likely 4-5 hours
combined into one file
10-min interval data from
Once at 6 synoptic sites
ProODO, min. 24 hours
during experiment (sites
can be combined in one
file)
10-min interval data from
Beginning and end at each
ProODO, upto 5 days
experimental site
continuous
experiment (sites can be
combined in one file,
suggest one beginning one
end file)
1-min interval data from
For all experimental reach
ProODO, a few hours
sites (three per year), can
encompassing ER and NEP be combine into one file
for all chambers
5-min interval data from
Once at 6 synoptic sites
ProODO, min. 30 min
during experiment (sites
before and after each
can be combined in one
deployment
file)
5-min interval data from
Beginning and end at each
ProODO, min. 30 min
experimental site (sites can
before and after each
be combined in one file,
deployment
suggest one beginning one
end file)
5-min interval data from
Once per chamber
ProODO, min. 30 min
measurements, three sites
before and after each
total per year, can be
deployment
combined to one year.
1-min interval data from
Once per biome and year.
ProODO, for as long as 1
mg L-1 change, and with
different light settings,
likely a few hours
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XXX999999_consumer_synoptic.csv Species, their length and
calculated weights for
synoptic sites
XXX999999_consumer_reach.csv
Species, their length and
calculated weights for
reaches of all experimental
sites
XXX999999_consumer_species.csv
Species list of all consumer
XXX999999_inverts_all.csv
Species, abundances, and
potentially lengths from
baskets of all experimental
sites (habitat and reach
scale)
Species list of all consumer
XXX999999_inverts_species.csv
XXX999999_ammuptake_reach.csv
XXX999999_ammuptake_patch.csv
XXX999999_waterchem_reach.csv
XXX999999_BOM_all.csv
XXX999999_chla_all.csv
Location identification,
timing of sampling in
chambers, and ammonium
concentrations
Location identification,
date, and different water
chemistry parameters
Location, date, BOM data
including Ashed mass,
AFDM, drymass for coarse
and fine
Location, date, chla data
including rock area, and
chla per area
58
Likely one file per year
with all three synoptic sites
listed
Likely a file for beginning
surveys at all sites, and an
end survey with an extra
file containing 10-d and
20-d control survey in
treatment reaches
New species when needed
additional files can be
created
Update whenever enough
data accumulated, likely
one per biome and year
New species when needed
At the beginning and end
of the experiment, for three
sites, likely combined for
one beginning and end file
Most likely one per biome
and year
Most likely one per biome
and year
For all experimental reach
sites (three per year), can
be combine into one file
For all experimental reach
sites (three per year), can
be combine into one file
SCALER methods Jan 9 2013 v 1.0
6.6.2. Raw data entry qa/qc
To avoid data entry issues spreadsheets with dropdown menus are available for all data collected under
the file names listed in table X.
6.7.
Backup procedures
All notebooks should be scanned on a weekly and the scans uploaded to the scanned notebooks
folder in the “data” folder of the “SCALER all” dropbox.
All data files that are to be incorporated into the database (i.e., all files listed in table X) need to
be placed in the new data folder in the “data” folder of the “SCALER all” dropbox. Raw data are then
incorporated into the database by the data manager and files moved to the “processed data” folder.
All data should be backed up on external harddrives as well.
7.
7.4.
Modeling procedures
Data calculation
Metabolism – chamber, 1-station, 2-station
Nutrient uptake – chamber, 1-station, 2-station
Aeration rates – 1-station, 2-station
Rock surface for chla
8.
7.5.
Workflow storage
7.6.
Input data file storage
7.7.
Output data storage
References
Benda et al, 2004
Bott, T. L. 1996. Primary productivity and community respiration. Methods in stream ecology.
Academic Press, San Diego, California:533-556.
Covino et al. 2011
Dodds, W. K., C. Randel and C. Edler. 1996. Microcosms for aquifer research: Application to
colonization of various sized particles by groundwater microorganisms. Groundwater 34:756-759.
Mulholland, P. J., C. S. Fellows, J. L. Tank, N. B. Grimm, J. R. Webster, S. K. Hamilton, E.
Marti, L. Ashkenas, W. B. Bowden, W. K. Dodds, W. H. McDowell, M. J. Paul, and B. J.
Peterson. 2001. Inter-biome comparison of factors controlling stream metabolism. Freshwater
Biology 46:1503-1517.
Riley and Dodds (in press)
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SCALER methods Jan 9 2013 v 1.0
Reichert et al. 2009
Tsivoglou and Neal (1976) gave best es
9.
Glossary
Biome – The five different areas participating in the SCALER project are referred to as biomes
to separate and avoid confusion with site (see below).
Site – Study area within a biome, either one of the 20 synoptic ones, or one of the 6 consumer
manipulation areas.
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SCALER methods Jan 9 2013 v 1.0
Appendix 1
Field material list for experiment at one biome in one year:
GPS
Electronic distance measure (meter tape as backup)
Meter stick
Wentworth scale
Multi-parameter meter
Spherical densitometer
Clinometer/hose
Measuring tape
Flow meter
Loeb sampler with brush (4.9 cm2 area) for bedrock and substrates with large planar area
Neoprene template (4.9 cm2 area) and toothbrush for smaller substrates
Small PVC cylinder (4.9 cm2 area) and non-slotted spatula for sampling soft substrates
White plastic or enamel tray
Whirlpacks or 1-L bottles
Squirt bottle for rinsing
Cylindrical sampling template (~30 cm diameter – i.e., LINX2-style “bottomless spaghetti pot”)
Sample bags for leaves and wood (10 per reach, 1 per sample)
Sample bottles for FBOM subsamples, approx. 250 mL size (10 per reach, 1 per sample)
Small plastic ruler
Rebar (~1 every 30 cm for 1.5 stream widths, for each of the three sites)
Exclosure wood frame: 19 in long wood pieces of 0.5x0.5in (WxH), screwed together in a square with a
20.5in piece stabled in the diagonal – 8 per experimental site, thus 24 total per year
Staple gun and staples (~200 per 8 exclosures, thus 600-700 per year)
Fencing (4ft high, 50ft long per experimental site, 150ft per year)
Fencing (3 or 4 ft high, 4 x 1.5 stream widths long, for each of the three sites)
Rebar (3 or 4 ft high, one every 30 cm for 4 x 1.5 stream widths long, for each of the three sites)
4 4ft ½ in rebar (8per experimental site, 36 per year)
Clippers
Zip ties (~300)
Mallet
Shovel
Note book, pencil
Backpack electroshocker
Dip nets
Buckets (20L, 1 marked by Liters)
Measuring boards
Seine
Minnow traps and bait
Waders
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Strawberry baskets (10 x 10 x 6 cm), 112 per experimental site, thus 336 per year
Mesh lining (if deemed necessary)
Bucket to collect and mix gravel
Bull’s eye level
Sample bottles (150 mL) 8 per experimental site and sampling time, thus a total of 96 bottles
7 chambers (bring one as reserve)
7 covers
Electrical boxes
12v battery
YSI ProODO meters (total of 20 available)
Odyssey irradiance meter (total of 20 available)
Tools (Allen wrenches, screw drivers, duck tape)
1L volumetric cylinder
Bucket with 8L mark
Bucket with 4 L mark
Funnel
Ammonium stock solution
Sample bottles for ammonium uptake (80 30 mL bottles)
Sample syringe and filter apparatus
Whirl packs (32), labeled
Cooler with ice
32 500 ml Nalgene bottles, labeled
White pan
250µm sieve (2)
Water squirt bottle
Forceps
Whirl packs (64 per experimental site, thus 252 per year)
Bottle with 100% formalin (diluted with stream water)
Field material lists by experimental day of each of the three experimental sites
within a biome for a year:
Day -1
Fencing (3 or 4 ft high, 4 x 1.5 stream widths long, for each of the three sites)
Clippers
Shovels
Rebar (~1 every 30 cm for 1.5 stream widths, for each of the three sites)
Mallet
Zip ties (about 3-4 per rebar)
Note book, pencil
Backpack electroshocker
Dip nets
Buckets
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Measuring boards
Seine
Minnow traps and bait
Waders
Day 0
Note book, pencil
Backpack electroshocker
Dip nets
Buckets
Measuring boards
Seine
Minnow traps and bait
Waders
Exclosure wood frame: 19 in long wood pieces of 0.5x0.5in (WxH), screwed together in a square with a
20.5in piece stabled in the diagonal – 8 per experimental site, thus 24 total per year
Staple gun and staples (~200 per 8 exclosures, thus 600-700 per year)
Fencing (3ft high, 50ft long per experimental site, 150ft per year)
Clippers
Zip ties (~100)
Rebar (8 4ft 1/2in)
Mallet
Shovel
Strawberry baskets (10 x 10 x 6 cm), 112 per experimental site, thus 336 per year
Mesh lining (if deemed necessary)
Stream gravel
Bucket to collect and mix gravel
Day 3
4 4ft ½ in rebar (per experimental site, 12 per year)
Mallet
4 YSI ProODO meters with extended battery compartment
4 Odyssey irradiance meters
Bull’s eye level
Zip ties
Sample bottles, labeled, 8 per experimental site and sampling time, thus a total of 96 bottles
Syringe and filter apparatus
Filters
Forceps
Cooler with ice
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Day 31
Nutrient uptake
Day 32
6 chambers (bring one as reserve)
6 covers
Electrical boxes
12v battery
6 YSI ProODO meters
Odyssey irradiance meter
Tools (Allen wrenches, screw drivers, duck tape)
1L volumetric cylinder, bucket with 8L mark
Funnel
Ammonium stock solution
Sample bottles (80)
Sample syringe and filter apparatus
Whirl packs (32), labeled
Cooler with ice
32 500 ml Nalgene bottles, labeled
Bucket with 4 L mark
100% Formalin
White pan
250µm sieve (2)
Water squirt bottle
Forceps
Whirl packs (32) labeled outside and in
Bottle with 100% formalin (diluted with stream water)
Day 33
Note book, pencil
Backpack electroshocker
Dip nets
Buckets
Measuring boards
Seine
Minnow traps and bait
Waders
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List of samples for 1 biome, 1 year
Water chemistry
68 (9 parameters proposed to measure)
Nutrient uptake
~850 based on 25 samples per pulse station plus chamber
samples (1 parameter)
SF6
~750 based on 25 samples per pulse station
Chlorophyll a
~200 (~100 from baskets, rest reach and synoptic)
BOM
~200 (~100 from baskets, rest reach and synoptic)
C:N:P
biofilm ~100 since reach and synoptic only; BOM ~300
since leaves, wood, and FBOM
Inverts
96
65