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Rithron Feeding Type Index is a German feeding index proposed by Schweder (1990). If I correctly understand it is calculated using the ratio:
(grazers + shredders) / (grazers + shredders + filter feeders + deposit feeders)
I have not found the traits table used by Schweder to calculate RETI (I do not have access to the publication) with the affinities of the different macroinvertebrates taxa for these 4 modalities (grazers, shredder, filter feeders and deposit feeder).
Does anyone have the traits table used by Schweder and could confirm that the RETI is calculated with this formula?
Many thanks in advance!
SCHWEDER, H. (1990): Rhithron-Ernährungstyp-Index (RETI) - ein Parameter zur Beschreibung und Bewertung der Ernährungsbeziehungen von Makroinvertetbraten in kleinen Fließgewässern.- Deutsche Gesellschaft für Limnologie (Hrsg.), Erw. Zu-sammenfassung d. Jahrestagung 1990 in Essen, S. 325 - 329.
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I found this pdf, hope to help you and it said based on individual numbers. Page 13 and 14
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I'm thinking of purchasing one and am interested to hear others experiences, especially problems they may have encountered.
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Hi Scott et al.,
We have had generally okay results with a benthotorch but only after we learned to be pretty constrained in how we apply it. One major issue that we found is that the benthotorch underestimates standing stocks on rocks that are taken from full sun. My grad student wrote a paper about this (A cautionary tale for in situ fluorometric measurement of stream chlorophyll a: Influences of light and periphyton biomass in Freshwater Science last year) in which they found that you needed to have the rocks held out of the stream for 15 to 30 minutes before you get reliable numbers. The company was pretty unpleasant to work with when we brought up this issue with them. There are also potential issues with self-shading when standing stocks are high such that you get underestimates of the Chl a. We have found that it works well on tiles and on NDS glass frits and that is where we use it most. As long as you pull your rocks out of the stream and cover them for a bit or if you do your sampling in shaded streams or early in the day before the streams have had full sun on them, they work okay for natural substrates. Overall, we found that being able to take a high number of replicates makes it a useful tool for comparison within and among sites/treatments and it's probably worth getting for that. But, it is not as good if you are relying on it for a highly accurate measure of Chl a.
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I have fish density data (n per 100m sq) for fish populations in three river systems. Each river has a site immediately above a effluent discharge, a site immediately below it, and two further downstream sites. Each site was surveyed biannually for two years. Is there a way to statistically compare densities between upstream and downstream sites that does not contravene the assumption that samples are from independent populations? I have evidence of limited migration of brown trout between sites on the same river. I have read papers that seem to ignore the assumption, and perform ANOVA or Kruskal-Willis between sites on a river regardless of the apparent pseudoreplication. Many thanks in advance.
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Hi,
To account for the apparent pseudoreplication on your river, you could just include it in your analysis via an asymmetric eigenvector map (AEM). This technique allows you to create directional eigenvector that correspond to your unidirectional phenomenon (here the river's flow). You'll only need your geographic coordinate if you use R. You can, then include those vector into the analysis of choise, as a new explanatory value and include the link between your site without ignoring the potential pseudoreplication.
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When looking at the effects of climate change on modeled streamflow, is the amplitude of the seasonal cycle (maximum month - minimum month, from an annual climatology) a useful metric? I can imagine this would matter for ecosystems who rely on streamflow velocities falling within a certain range, or for reservoir operations in general, but I'm struggling to find articles discussing it. Does anyone know of a good source addressing the importance of seasonal amplitude? Or is this not an important consideration?
Thank you!
Meg
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Dear Megan,
the importance of the seasonal amplitude of streamflow is enormous from an ecological standpoint in many systems, e.g. in non-perennial rivers and streams (NPRS) (see e.g. Skoulikidis et al. 2017 and references therein). This was even one of the main points determining the evolution of the concepts of ecologically sustainable flows from a minimum vital flow to environmental or ecological flows since the 1990s.
Related to this is the ecological relevance of the spatial and SEASONAL extent of Water-Level Fluctuations (WLF) in lakes and reservoirs (e.g. Leira & Cantonati 2008, attached) and of level drawdowns in rivers.
Another example is the large relevance of seasonal variability of discharge in special habitats such as springs that are often biodiversity hotspots in spite of typically small size (see e.g. Cantonati et al. 2012 attached).
Best wishes.
Marco
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Dear friends,
I am submitting a manuscript with following highlights. Would you please take look to them and give me your opinion?
· Energy flux and mixing process mechanistic model, i-Tree Cool River, was assessed in steady and unsteady conditions.
· The warming effect of urban storm sewer on the river during storm events.
· The cooling effect of riparian vegetation shading and subsurface inflow in the summertime.
· Linear interpolation, Gaussian Elimination function, in C++ for matrix operation.
Best,
Reza
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Dear Reza, in the two middle sentences I am afraid there's something missing, a verb? Probably can end more easily readable if you reformulate them. For example,
Urban storm sewer had a warming effect on rivers water during storm events.
Riparian vegetation had a cooling effect.....
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Identification, please what is it? I can't identify. it is from freshwater (stream)
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Hi,
Daniel is right, it is Tabellaria flocculosa. The "zig-zag" bands are the result of divisions - one pole is still attached to the next cell via extracellular poysaccharides, the other not. This is the so.called side- or girdleband-view.
Regards
Michael
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Hi everyone, I am currently working on my Master thesis about connectivity of native riparian forests in Uruguay. For that reason, I need the (mean) flow velocity [m/s] of Uruguayan rivers to calculate dispersal ranges via hydrochory for some woody species.
Do you know if this kind of data already exist? And/Or who I might ask to get these information?
Thank you.
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other papers attached. I hope they can be useful or you can contact corresponding authors to have more information.
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For my MS thesis I am researching how flow reduction will affect macroinvertebrate populations in a river system in Utah. Over the past fifty years, water management agencies have sent high volumes of water through this river, with profound implications for its aquatic ecosystem. Now they are wanting to reduce the minimum flows. I will be collecting macroinvertebrate samples along the length of the river for two years: the first year being an artificially high flow regime, the second being a lowered, more natural flow regime. With discharge as my independent variable, I hope to observe any trends in Functional Feeding Group abundances / biomasses over time, as well as trends in the sensitive / tolerant taxa ratios.
Since I'm less concerned with indices of diversity, would identifying macroinvertebrates to family be sufficient? My concern is that since different genera within families may fall into different FFG's, this taxonomic resolution might not accurately reveal the patterns. Are there any methods of data analysis that I could employ to work around this? Or - knowing that this study limited in both time and money - what families would you recommend I take down to sub-family or genera, if I were to choose only a few? 
Thanks in advance!
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I agree with Timur and Kevin (and others).  You'll pick it up rapidly.  Your first couple of samples may seem challenging, but it will get incrementally better from there.  You'll rapidly build up a list of the "usual suspects". 
Some considerations:  1. Chironomidae is THE common dipteran family that will probably stay at family unless you invest significant time and effort mounting heads (or find a consultant if you can afford one).  2. Getting the EPT families down to genus is well worth it; they respond to impacts. 3. If your samples are quite large then consider sub sampling with a goal of around 300 macroinvertebrates per sample (Look up Vinson and Hawkins work supporting that number).  4.  Stratify your ID work....ID your sampling sites and sampling times in rotation building up your replication in each treatment as you go.  If differences are large then you will detect them with few samples and you can then decide how many replicates you truly need to ID (it's usually fewer than you think). 5. The functional feeding group indices may not be the best way to detect differences....diversity indices that include the EPT taxa may be more effective.  6.  Contact your Utah DNR folks and get a list of what they find....especially if they work your river...this will help with your taxonomy.  7.  If you are permitted to hire undergrad workstudy kids....do it....and be prepared to work with them.  They can pick, sort, and if you get reliable ones they can ID. 8.  Have fun with it!
Here's a paper I did with undergrads.....generally to genus except the midges.  We evaluated several indices by building up replication level until we were able to detect differences: http://academics.smcvt.edu/dmccabe/research/PubsPDFs/Standardized%20effect%20size11-080.pdf
Cheers
Declan
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I am curious as to whether there is any literature on how in-stream coarse woody debris levels may change in relation changes in species composition.  In particular I am working in a system where trees are frequently "caught" in an invasive woody shrub.  
It is easy to measure dead wood on the ground, or standing (snag), but what about actually caught in other live material?
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Are you intend to measure each individual CWD? How about measuring the size (diameter at both ends and length) and estimate the density (using penetrometer or decay class)
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I found this green water in a gold-mining basin collecting rainwater in French Guiana where freshwaters are normally dark-brown to black coloured due to the humic acids. I suspect blooming of cyanobacteria. Can anyone have any suggestions to explain this unusual colour?
Kindly help me in this regard
Thank you
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Be careful if sampling, some of the blue green algae can be hazardous or toxic.  Personal protective gear may be needed and best to avoid contact until you know what it is.  Blue green algae can fix their own oxygen and CO2, so if the temperature and alkalinity/pH systems are out of balance, green algae cannot live and blue green develop.  I dont remember explanation for specific color, but you can look that up on the internet.  Also I dont know if they are using mercury or other chemicals to concentrate the gold, but if they are using hazardous materials, they may have altered the conditions for algae.  If looks like this is settling ponds, so the water is not being circulated or diluted, so the temperatures and nutrients are elevated, and open to sunlight also encourages algae growth.  Please dont let them dump this in the river, it will probably eventually go away.  If you test water quality for alkalinity and pH, if your alkalinity is low and/or pH high, blue green algae may invade.  It has been a long time since I studied this, so best to consult an expert and make sure you have the proper permissions if treating.  Hopefully this will help.  
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hi. im working on abundance of macroinvertebrates in seagrass community. i am using a 2D MDS to see their resemblances, which are at 60 and 80 percent. however, there are points that deviate from the groups. how should i interpret this? thanks! :D
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Hi,
as you my know the MDS is a graphical representation of the similarity matrix, closer points means that these are more similar than some that are further apart. If I get your description right the points that clearly deviate from major clusters are below the threshold of 60% similarity.
There are various ways to see how variation is distributed in your data such as Principal Component Analysis (PCA), ANOSIM, Principal component Analysis (PCO) which may help you to interpret the reasons for the underlying grouping.
Cheers
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I am interested in evaluating the potential dispersal of mayflies nymphs in streams to quantify dispersal as a structuring mechanism of metacommunities. I have tried some available dyes (such as rose bengal), but they are not being efficient.
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Did rose bengal kill the mayflies? Or was it simply not taken up by them?
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Good Day,
I am trying to figure the analysis I need to do for my project. I study shrimp and fish in 3 tropical intermittent streams. I am testing to see if there is a difference in species composition at different elevations. My experimental design includes a low, mid-, and high elevation site in each stream. I have selected 3 streams, so therefore 9 sites total. I chose 3 streams so I can say with confidence that a species distribution changes with elevation. I have been sampling for a year. I have sampled at each site 4 random times in the year; therefore, I have collected 12 samples for each stream (4 at the low site, 4 at the mid-site and 4 at the high elevation site). Altogether 36 samples. We have a dry and wet season here. Seasonality is a covariable.
What is the best way to setup my data? Recommendations for analysis? Help is much appreciated:)
Thank You,
Kayla
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You are doing sort of a repeated measures analysis of variance, or, as mentioned above, repeated measures analysis of covariance. In JMP, the organization of the dataset has to be the way JMP wants it (which requires a little manipulation from the most sensible way to input the data) and you have to have sample ALL moments in ALL treatments. If you have empty cells, it makes it more complicated, although there are ways to get around it. It is beyond the limits of this medium of discussion to explain how to rearrange your data - although the help in JMP is pretty good at explaining it. Also, your sample size is very small for such a complex analysis, as Maurizio suggested. In general, a multivariate scheme requires an N of 25 plus 3 times the number of treatments as a minimum. If it is very complex, add 25 per level.
I hope this helps!  Jim
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How to determine the daily streamflow of a river while fully accounting for climate change aprt from using hydrologic models such as SWAT, WEAP etc? Can the ARIMA models be twigged to suit this purpose?
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SWAT and/or Hydrological models are different from ARIMA. ARIMA needs only observed data............. but SWAT requires too much info including observed data for calibration and validation. The challenge is to obtain all info for physical-based Hydrologic models
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Hello!  I am looking for recent (ca. 2005-) literature on the (typically beneficial) effects of riparian buffer zones on receiving freshwater ecosystems, particularly lakes, streams, and groundwater.  Reviews and papers on empirical investigations will be equally welcome.  Thanks a lot in advance!
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Hi Paola,
This featured collection on riparian ecosystems and buffers might have some useful references - see Mayer et al (2014) for intro: http://onlinelibrary.wiley.com/doi/10.1111/jawr.12212/abstract
Also for a review on the beneficial effects of riparian vegetation on water quality see Dosskey et al (2010): http://onlinelibrary.wiley.com/doi/10.1111/j.1752-1688.2010.00419.x/abstract
Hope this helps.
Marian
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According to my paper collection the SASS (South African Scoring System) method is the standard for the bioassessment of rivers in South Africa. The SASS was adapted to be used in other regions of southern Africa, including Zimbabwe Swaziland, Namibia (Namibian Scoring System NASS), Botswana (Okavango Assessment System OKAS), and Zambia (Zambia Invertebrate Scoring System ZISS). Recently, one paper and two master theses confirmed that the SASS approach does basically work also in other parts of the black continent, namely Ethiopia, Uganda and Burkina Faso. I would like to know if the SASS methodology is used in more countries than I listed above. By "used" I mean for example adopted (directly used), adapted (exclusion or inclusion of indicator taxa, change of rankings etc.), or used as a metric in a multimetric procedure.
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Otto, in Tanzania also there is a development in the use of SASS in a modified version called TARISS (Tanzania River Scoring System).
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Landscape commuity genomics is related to eco-evolutionary processes in complex environments, such as stream and riparian ecosystems. However, its framework is not clear at the moment, because we don't know how genomic variation is affected by dynamic interactions between abiotic (environmental) and biotic (community) effects..
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Thanks to Francis for linking in Hand et al.'s essay. Very interesting, and one of those concepts that's obvious...after it's pointed out. As Edward Tufte reminds us "It's More Complicated Than That!". The research implications? As usual, it's a case of a big idea that needs to be translated into testable hypotheses on scales that can be studied on practical levels. In riparian corridors, you might expect -for example- genetic variation to be more highly correlated along the corridor, where conditions are similar, than across a stream-to-upland gradient. Or maybe the opposite, depending on the organism.
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Wood, leaves, algal clumps and other particulate organic matter (POM) enter the Great Lakes from streams that flow into it.  I'm curious how important these inputs might be to lake carbon budgets.  Has anyone quantified stream POM inputs or modeled this?  I'm especially interested in Lake Superior. 
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First of all, I measured the DOM from stream to lake ecosystem, but not the POM. Of course, they play important role in organic matter bugdet in lakes. You can read my paper from the reference (Mostofa KMG, Yoshioka T, Konohira E, Tanoue E, Hayakawa K, Takahashi M (2005b) Three-dimensional fluorescence as a tool for investigating the dynamics of dissolved organic matter in the Lake Biwa watershed Limnology 6: 101-115).
POM play an important, specifically depending on the contents of different organic matters (wood, leaves, algae, etc) and subsequent decomposition in lake bed and surface. For example, alage can release DOM more quickly, but wood may release decompose for weeks to months in lake bed. This should be considered clearly. You can also read book "Photobiogeochemistry of Organic Matter: Principles and Practices in Water Environments" that publsihed by Springer in 2013.
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I'm trying to calculate secondary production of freshwater benthic communities. Some times (nearly in 50% of cases) I got negative numbers. Does it means that destruction in bigger that production?
I'm want to understand if I have done the mistake or not.
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@Irene: I reiterate that the formula does not work.
When you use B to estimate P (the P/B method), predator consumption is already accounted for (they ate some of the B of the non-predators). So the production available to fish is the sum of the production of predators and non-predators.
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We use a short range DIDSON in Redwood Cr, and a long range ARIS in the Mad R. Redd surveys are independently counted in Redwood Cr. To date (3 yrs of data), we are not seeing any agreement with sonar counts and redd counts for Chinook salmon and steelhead trout. The redd counts are negatively biased, even if you take into account that not all salmon passing the sonar beam will spawn (pre-spawn mortality). I know there is similar work on the Secesh R in Idaho, and I am wondering if any other place has a similar study going on and what their results might be.
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We contract annual redd surveys of spring Chinook salmon through Oregon Department of Fish and Wildlife. I PM'd you the POC as well as a DRAFT report. Typically to estimate the escapement for a reach the redd count is multiplied by 2.5. This conforms to the negative bias you are seeing. However, we don't have DIDSON monitoring, and instead use the video counts at barrier dams.
What is interesting is that the productivity and effective population size of fish transported above our projects can vary widely by watershed . We are monitoring this through parental based genetic studies. In one basin we transport over 500 adults yet the number of individuals contributing to the cohort for that brood year number lass than 100. This is likely due to a combination of PSM and losses during incubation. It is likely that the interbasin variation is likely due to the hydrological and geological differences. Our lowest performing subbasin has the combination of large floods during incubation, combined with a river channel that has a lot of bedrock.
We also have done a lot of investigations into pre-spawn mortality which I can provide if it would be useful.
v/r,
Griff
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Could vegetated gravel bars be classified as riparian zones?
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This is more a matter of terminology than a discussion on the role of those vegetated gravel bars within the river landscape. The riparian vegetation is the herbaceous or woody vegetation located at the interface of two ecosystems - the terrestrial and the aquatic - and it is not relevant for this that the river is chanelised or not. However, if those gravel bars are inside the riverbed and subjected to submersion caused by floods, that vegetation should not be considered riparian. So, you have to figure out if that vegetation is located in the in-stream channel, inner banks or outer banks, to name it correctly.
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Is anyone currently examining stream grazer foraging strategies in terms of how they partition periphytic resources?
In particular, are there any trait-based models (or ideas) out there predicting how stream grazers will forage across larger scales (e.g. meters)?
There is the classic Steinman model of small-scale (micrometer) utilization based on periphytic structure, but I'm trying to think "big" here.
I welcome people's thoughts & ideas.
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Check out the publications in the Researchgate profile of Jan J. Verspoor. He looked at salmon interactions with stream periphyton at  watershed scales - might be some useful resources in his work.
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Petit test, Double mass curve technique, Mann Kendal Rank Test etc...which is more hydrologically significant technique for change point detection. Please help me.
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Different approaches have been found to be effective for identifying different types of changes so the answer will partly depend on the nature of changes that you suspect may be an issue for your data set and the type of changes that exist in the data set.  Hence, applying many of the techniques suggested so far will be useful to help learn more about your data.  One additional technique to consider is Bayesian Change Point Analysis, which is available through the BCP package in R.  For any changes that are found, it is important to examine the metadata for your time series to look for any changes in measurement protocols or equipment that may explain the changes as well as any known physical changes that could explain the change (i.e., a forest fire on a watershed can result in changes to the streamflow time series).
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I have a stream where the seasonal maxima oxygen concentrations (i.e. annual max of daily max) preceded measured abundance peak by a few weeks. Of course the abundance measure has it's error. But please ignore that for now and think about whether oxygen production should, in theory peak before, at, of after plant maxima. A plant biomass peak defines the transition from net growth to net decline. Would it be safe to assume net growth is levelling off prior to the peak? Would oxygen production reflect biomass of total plants, biomass only of the growing cells, or the net growth of plant material (e.g. kg dry matter added per square meter per day). The stream oxygen measurements reflect what is happening over the reach upstream on a given day, so think about reach scale biomass/growth as the sum total of individual cells within individual stems. 
Thoughts and suggestions welcome, thank you.
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Good points - thank you.
I have estimates of photosynthesis (g/m3/d at 20C). Reaeration calculated from time lag between solar noon and DOmax. Respiration and photosynthesis calculated using delta method. Is it reasonable to treat that photosynthetic rate as net prim. productivity?
This gives a similar result with max photosynthesis preceding max plant abundance by about two weeks. I get a similar lag with depth-corrected photosynthesis (g/m2/d at 20C, using a time-series of depth), and with P/R and P+R. Within a few days of each other. Probably worth mentioniong that the end of season decline in plant abundance had the most pronounced lag after the decline in photosynthesis (closer to 4 weeks).
Of course I don't have a true measure of plant abundance, and I can write the lag off as an artefact of the methods I used to estimate abundance (%bed cover). If I could measure true population biomass, would that determine net primary productivity given that not all of the plant biomass is actively photosyntheisizing (e.g. shaded, structural or sensecing material)?
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I'm interesting in linking several fish habitat sample protocols over a long time period to describe nearly a century of change. Bureau of Fisheries reports from the 1930s, Aquatic Habitat Inventories from the 1990s, and the Columbia Habitat Program at the present. The general problem is one of describing change in a consistent manner although protocols have changed drastically and there may be no or little crosswalk between the methodologies. Any suggestions for methods or relevant research would be helpful. Thanks.
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Hi Seth!
Some kind of 'reference condition approach' could maybe work. This is usually used for fish or invert assemblages, where you set a reference condition based on undisturbed streams and then calibrate your metric based on that reference condition. You will always get a score between 0 and 1, which can then be compared to a a different measurement system. There's a few papers on this flying around, including one I was involved in where we modeled pre-european habitat conditions in Australia,
Cheers
Simon
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How do I use HYDATA? What type of data does the model use? Are there manuals whether pdf/word/video tutorials?
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I have hundreds of small sources which have many sinks at a local level and then together drains towards a main sink at the end of the stream. How do I define it in ESRI ArcGIS?
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In this, create a vector data after that you may try to analysis it, the following link may useful for you. http://resources.arcgis.com/en/help/main/10.1/index.html#//002r0000002m000000
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There are a number of studies which show little or no effect on invertebrate communities, but these may have been affected by the absence of source populations and dispersal-limitation (e.g. Sundermann et al. 2011 Ecol Apps), or potentially, spatial and temporal legacy effects. I'm interested to hear about studies that showed positive effects on stream invertebrate communities (e.g. increased abundances of EPT) through habitat augmentation and rehabilitation.
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See the US Army Corps of Engineers studies of Nine Mile Run in Pittsburgh Pennsylvania. An urban stream with a range of complications that was radically reconfigured with a new stream bed and riffle pool sequence designed to withstand the challenges of urban stormwater. Each year a benthic study and fish study has been done with significant positive gains over time. Mike Koryak was the original stream ecologist with the US Army Corp of Engineeers, now retired. Mary Kostalos did benthic studies in feeder creeks in Frick Park as well as in NMR throughout her career at Chatham College and may be still doing work in the field there.
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I have 100ml of water containing some organic particles <1mm in diameter. I want to extract and weigh all of the FPOM, but am struggling to find a cost-effective way to do this. Has anyone done similar work and can advise on the best technique? (I'm leaving the definition of FPOM deliberately ambiguous as this seems to vary wildly in the literature - feel free to discuss this in your answers too).
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Cost -effective makes it tough. Silver 0.45 micron filters and a 100 ml syringe would be best e.g. from Millipore. Clean the filters by ashing, use then to filter your water, measure TC on the filter by combustion.
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I would like to analyze a test of aquatic insects, indicator species of headwaters.
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Hi Hugo,
You could use the R package labdsv for running IndVal: http://cran.r-project.org/web/packages/labdsv/labdsv.pdf .
Best wishes,
Jani
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Limited material preferable, power equipment unfortunately not a viable option
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You could potentially use over the stream to induce a greenhouse, causing warming. However, you would probably need a pretty long reach of this warming setup as the water is (obviously) continually renewed. Many terrestrial climate change studies use plexiglass roofs to artificially increase temperature, but they don't have to deal with running water.
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For i.e. EPT index, ASPT, BMWP, etc.
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Hi Rosanna,
Sure thing will email the information to you tomorrow. I assume you are aware of the BMWP scoring system primarily used for detecting organic pollution events. One current shortfall in this method is that it fails to utilise abundances, although this soon will be superseded by the WHPT index which will have pressure sensitivity scores for each scoring family linked to abundance categories.
Kind regards,
Drew.
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The sediment and organic matter caught with the biological content of the kick sample, I'm guessing will vary somewhat with operator and technique. I am collecting habitat data separately for the areas kick sampled, but would the material in the sample act as a useful proxy for habitat? I might just try it anyway since I have the material in the store.
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I can suggest you to investigate your sediments collecting them directly (it is already an approximation, without taking into account the variability of the operators).
In our lab we evaluate the dry weight of about 10 grams of sediments (48h 60° C)
The dry sediment is collected in crucibles (ours are very small, less than 1 gram capacity) and treated for 8h at 450 ° C to obtain the amount of organic matter (for difference in weight). With larger crucibles (up to 10 grams) the time lengthens to 12h.
For the inorganic matter we process samples for other 8h (or 12h) to 900 ° C.
For the particle size analysis you need more material, 200 grams dry weight is a good amount for our tools. If you are sediments of freshwater environments must first be treated with H2O2. I uploaded two jobs on my profile, unfortunately, are in Italian (Protocollo di campionamento ed analisi chimica di sedimenti lacustri Protocollo di campionamento ed analisi granulometrica di sedimenti lacustri). If you need more information contact me .. I hope to be helpful
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Entry of organic contamination through fecal debris of animals and leachate from the adjoining reserved forest cover area during rainy periods.
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Answer
I generally agree with the above mentioned theoretical consideration that BOD is the part of COD that can be oxidized biologically by microorganisms.
However in practice the two parameters are defined by analytic procedures. If one uses standard methods with an inocloum for the BOD test of activated sludge and the COD is determined by the oxidation of chromate it happens in both wastewater and polluted streams that the measured BOD is higher than the COD because chromic acid does not oxidise ammonia while a bacteria community with nitrifiers does this quite efficiently.