Michael F. Merigliano added an answer:What method should I use for comparing similarity of species compositions among subsampled sites?
I have sampled 4 sites for bryophytes using quadrats in and around 4 wetlands. (I recorded abundance data.) Two of the wetlands are embedded in deciduous woodland, two in coniferous woodland.
I'm trying to ask two main questions:
-whether the species compositions of the wetlands are more similar to the other wetlands or their surrounding woodlands
-whether the species compositions of the wetlands in each forest type are more similar to the other wetlands of the same type or to a wetland of the other forest type
Is there a test I can do to answer these questions?
I've done an NMDS ordination of the data. The polygons surround all the quadrats from each of the four sites, the spiders connect the wetland and woodland quadrats from each site. The wetland quadrats are colored blue, woodland green. Conifer is the plus hardwood is the triangle.
It seems to show that the two largest wetlands are most similar to each other (despite being in different forest types). It seems to show that the smaller wetlands each are more similar to the larger wetland of the same forest type. (Although the hardwood wetland is almost as similar to the surrounding woodlands as the other hardwood wetland.)
Is there some way to test if this is actually significant?
Although somewhat off-topic, a concern is the very small sample size (n =4, at most). There may be enough power if the differences between sites are very high with little noise, but your NMDS graph indicates otherwise.
A good, basic source for metrics and analysis of community data is McCune and Grace 2002. Analysis of Ecological Communities.Following
Valério De Patta Pillar added an answer:How can I compare community functional diversity across space and time?
I am working with a dataset of six communities that were sampled at two time points. I am looking for a simple way to see whether functional diversity of each community has changed from time point A to time point B.
My functional data is count data [number of species per community with a given functional trait]. As such, my dataset seems too small for most ordination approaches. Overall, I am investigating multiple traits within three different functional categories [growth form, habitat preference, and symbiont status]. At the moment, I am treating each functional category as a separate dataset.
Dear Klara, it is important to clarify your question:
You could compute functional diversity of each community at time point A and compare to functional diversity of the same communities at time point B. For this I suggest using Rao entropy, for which you will need to compute a dissimilarity matrix between species based on their traits (if the traits are of mixed type, the Gower index may be useful). In this case you can only tell about the temporal change in the overall functional diversity of each community, and nothing about the functional identity of the community components. This is a limitation analogous to when communities are compared by their species diversity (e.g., by using Shannon diversity).
Another option is to compare the communities based on their functional composition, for which you may consider the definition of fuzzy-weighted community composition. This is described in Pillar et al. (2009, http://dx.doi.org/10.1111/j.1654-1103.2009.05666.x). See also Pillar & Duarte (2010, http://dx.doi.org/10.1111/j.1461-0248.2010.01456.x) in the context of phylogenetic analysis. The fuzzy-weighting requires as input a similarity matrix between species based on their traits (which could be the above mentioned Gower index), which after proper standardisation to unit column will define a fuzzy set matrix U of species by species. The fuzzy-weighted community composition is given by matrix X obtained by matrix multiplication (X = UW), where W is the matrix of species composition in the communities.Following
Bernabé Moreno added an answer:Which biotic & abiotic components should I sample/measure in order to elucidate if a system is rather pelagic- or benthic- controlled?
Benthic realm and its biotic composition is highly important in glacio-marine fjords, specially when considering the cryosphere dynamics and the resulting phenomena (sedimentation, resuspension, freshwater influx, inter alia).
Also, in Antarctic seasonal bays (i.e. Mackellar Inlet (King George Island, South Shetlands), where I've sampled macrobenthic communities for previous research (see: http://goo.gl/YOy16D)) pelagic realm also plays a key role in terms of primary production and its consequent influence on higher trophic levels.
It's certain that analysing the benthic composition is more predictive when trying to speculate future scenarios. I presumably assume that the Mackellar Inlet is mainly a benthic-controlled system. Nevertheless, in order to be sure of this hypothesis I should go further through an integrated analysis of both realms.
The protocol that my colleagues usually execute is: macrobenthic survey (van Veen grab 0.05m2), collect plankton with plankton nets, and measure abiotic variables like temperature, pH, conductivity, turbidity and marine currents patterns (speed & direction).
I'd like to know if there's any specific protocol pointing straightforward to my question. What other measurements should I consider?
Thanks for the help. Cheers.
∞BSc. Bernabé Moreno
Thank you all for your substantive comments. They're already being very useful to help me structure some kind of study. Cheers.Following
Joe Shelnutt added an answer:Does anyone have suggestions for developing a course ecological recovery potential screening tool, to address multiple ecosystems across a landscape?
The location is a corridor from the Florida panhandle, westward to east Texas. Longleaf pine is a significant component of this landscape but riparian, wetland, seepage bogs, and other fragmented ecosystems need to be addressed simultaneously within this landscape. Developing a course tool to prioritize connectivity and where ecological restoration should/can occur is needed.
Thank you Olga and Joanna! I appreciate the suggestions and tools very much. Please let me know if I can help you. JoeFollowing
Mika Yasuda added an answer:Is functional alpha diversity equal to functional richness?
Pool et al (2014) quantify alpha functional diversity as the volume of the convex hull filled by the fish species of each community in two-dimensional functional space using the values from the first two functional axes.
But I wonder taxonomic alpha diversity is simply the species richness, so the alpha functional diversity can be functional richness...
Thank you very much for your answer, Thiago,
I will read these paperes as well.
Alan Feest added an answer:Should Simpson's E or Shannon's E be preferred for evenness estimates?
I have a multi-year dataset with about 100 species where the Simpson's D is already available.
As I want to look at evenness I wonder about the differences between Simpson's E and Shannon's E, and if any one of them is to be preferred.
I agree that a single measure of biodiversity is not very informative and my work on "biodiversity quality" is based on the picture expressed by a range of indices. Simpson's E sometimes shows change first when biodiversity is changing as it reflects the relationship between species populations.
Hope this is helpful.
Sebastian Dardanelli added an answer:Is it possible to incorporate detection probabilities to nestedness analysis?Is it possible to incorporate detection probabilities to nestedness analysis? The only approach I have seen that specifically deal with this issue are the papers from Cam et al. 2000 (Ecol Appl. & Oikos).
Hola Alexis! Si lo tenía a ese gracias! Parece que además de esos papers de Cam y el weighted nestedness no hay nada más. Voy a probar con esas dos cosas juntas para presencia ausencia y presencia ponderada por abundancias.
Irene Zweimüller added an answer:Which clustering method is the most realistic and descriptive in ecological terms?
I know that both 'Single-linkage' and 'Complete-linkage' clustering are monotonic methods (non-metric), which is great under a theoretical point of view. Nevertheless, 'Group-average' clusters sometimes are easier to describe.
I'm working with Antarctic macrobenthic communities, and I would like to know which of these three methods is the most adequate for an environmental analysis based on your own experience.
Thanks & cheers!
When you use cluster analysis, I recommend to standardize your variables (for example z-transformation, which results in mean=0 and SD=1 for the z.transformed variables). Ward´s clustering method (together with squared Euclidian distances) gives the best results - at least in my experience.
If you have extreme values in your variables, you may need to do a log-transformation before you start.
good luck, IreneFollowing
Raymond K. Timm added an answer:Is it worthwhile to only report a pattern in community ecology, without including an analysis of the potential environmental variables explaining it?
I was recently told by a referee that my manuscript about the long term temporal patterns of a fish community in a tropical wetland was irrelevant and useless because it did not include an analysis of the potential factors explaining such pattern, meaning environmental variables. These were not included because information for the years of my study is not available.
I would like to know if this is a widespread opinion among community ecologists, and if the state of the art is such that works limited to describe community patterns are not necessary anymore.
Thank you, I am looking forward to get to know your opinions.
Gabriella - sorry to hear about the harsh review. Nobody deserves that. I would chime in to say that it sounds like your study is stuck in the landscape ecology hinterlands between descriptive and functional metrics. in landscape ecology, functional metrics are of course the objective. With that said, describing patterns quantitatively is an important part of understanding how landscapes function - and has to be done first. If you don't already have it, check out FRAGSTATS and associated publications (Kevin McGarigal). They developed a free and very powerful utility to quantify patterns. Also, I might suggest resubmitting to Landscape Ecology and do what you can with other environmental response data to perform analyses and frame up hypotheses to build on your progress. Good Luck.Following
Elizabeth Kierepka added an answer:What is the best way to import microsatellite data to perform a dbRDA?
I am interesting in looking at the relationship of several environmental variables on genetic differentiation across 16 bee populations (Many individuals per population) using dbRDA. Using Legendre and Legendre (2012) as a guide, I have generated a distance matrix for my microsatellite data, and then performed a principal coordinates analysis. I now have two questions:
1. Negative eigenvectors need to be a corrected. Which would be the best method to do so?
2. The PCoA gives me an eigenvalue for each population at each axis, as well as the overall eigenvalue for each axis. I am assuming that it is the eigenvalues for population at each axis that are important. Is there a standard format used by vegan for this type of data?
I am new to analyzing community-type data, so any suggestions of pointers would be greatly appreciated. Thanks!
For dbRDA, there are R packages and separate programs that will perform the dbRDA all in one function. I have used vegan with the function capscale as well as DistLM by Anderson 2003 (PC program). Typically in a dbRDA, all the axes with positive eigenvalues are retained (at least in the calculation methods I have used), and axes with negative eigenvalues are not used in the analyses.
What you will use as your dependent variable are the scores from the PCoA, but any dbRDA function will do that for you. I also suggest that you run a partial dbRDA because you will have issues with spatial autocorrelation if you are running any community or spatial type data in your analyses.
I have code on Dryad from my previous manuscript comparing dbRDA and others in landscape genetics, but briefly here's what I did in vegan:
###gen file must not have headers, and it needs to be either a square or triangular matrix#######
This will give you your F-stats, inertia, and p-value for each landscape variable you tested. If you want the F-stat, inertia, and p-value for the full model, just leave off the by="term".
Hope this helps,
Sergey Popov added an answer:What software can be used to study the structure of a metacommunity?
Dear colleagues, prompt software for calculation of parameters a metacommunity. It is desirable to be able to use tables Excel. How can I adequately describe the effect of spatial scale on the part of the community?
Вот консольный режим и непривычен. Но, видимо, все равно придется и его освоить. Пока у меня основные программы SPSS, MapInfo, QuantumGIS, Surfer и серия программ для методов SADIE.Following
Summer M Burdick added an answer:Which is the best method to relate species occurrences to human impact variables in a sparse data matrix?
I have a data-set of macrophyte species (Presence/Absence, %-Cover) collected in river sites together with environmental (e.g. discharge, river slope, substratum composition...) and human impact variables (e.g. morphological quality scores, %-landuse in catchment...). The species data matrix is rather sparse, meaning many species occur in a few sites but are absent from many others, also because the data has been sampled along a long environmental gradient. I now want to quantify how species occurrence to particular environmental variables (either single or jointly) and particularily to human impact. One tricky thing in this regard is that some of the human impact variables are covarying with important environmental variables (e.g. discharge ~ %-landuse), making it challengig to quantify the effect of humans on species occurrences alone.
I already came along a number of techniques used in this context (e.g. CCA, INDVAL). However, I still concluded this is an interesting subject to discuss and hence I would like to hear your personal opinion about i.) the kind of analyses you would consider most suitable and ii.) how to deal with the sparcity of species and the mentioned covariation in the data-set.
Thanks, and I look forward to hear from you!
You might consider using an occupancy modeling approach (MacKenzie et al. 2006). This approach is good when your response variable is binary (presence/absence). There are occupancy models that allow you to look at the presence and absence of multiple species along an environmental gradient. One benefit of this approach is that it allows you to compare specific competing hypotheses.
You can deal with correlated parameters in several ways. 1) Where you have two variables just pick one 2) you could use a PCA or some other method to distill these variables into one meaningful component that can be used in your occupancy models or 3) you could fit a series of occupancy models that include and interaction between correlated terms. If you then compare these models with AIC you may be able to determine which variable is more important. However, if you have completely correlated variables you may never be able to tell which one is causing and effect.
I hope this is helpful.
MacKenzie, D., Nichols, J., Royle, J.A., Pollock, K., Bailey, L., and Hines, J., 2006. Occupancy Estimation and Modeling: Inferring Patterns and Dynamics of Species Occurrence. Elsevier Academic PressFollowing
Ken Sulak added an answer:Is it necessary to standardize for sampling effort before comparing asymptotic species richness?
If you want to compare diversity e.g. Hill numbers across species assemblages you need to standardize for sample size or sample completeness. Does the same count for asymptotic species richness estimators (such as Chao indices or ICE and ACE)? You calculate the same, namely 'true' species richness (often based on the number of singletons), so I guess it already corrects for differences in sampling effort?
Second (related) question:
Can anyone explain why I find different values for the same estimators calculated with different programs? Using the same abundance dataset I calculated ACE and Chao1 in EstimateS, R (fossil) and SPADE...and I got three different values for each estimator.
I know that EstimateS uses resampling and that it cannot calculate ACE, but strangely the Chao1 values are lower than the estimated species richness when doubling the reference sample.
None of the conventional measures of diversity and species richness, like H and H', are truly sample size independent. They are advertised as such, but fail when sample sizes are low. Species rarefaction curve analysis is perhaps the best way to compare species richness - if you have a number of samples from each study area to enable rarefaction curve analysis. There is excellent, inexpensive, and easily usable software to apply to the analysis. This is EstimateS. I recommend it very strongly. Easy to use, can dump Excel or flat data files straight in, and 95% confidence bounds are calculated. The fundamental rationale is that a large number of random draws are made upon the grouped data for each comparative study site. When the ESPP curve becomes asymptotic at X species, you can determine a Y number of pooled specimens that provides a solid estimate of species richness at that group sample size - for comparison with other pooled samples from other sites. You can plot the 95%CIs around the curves, evaluate for overlap. If not overlap at the selected Y number of specimens, then the two sites are significantly different. There are other softwares out there to do the same thing - but none is so easy to use.Following
Giovanni Gaglianone added an answer:Is there any publication on scientific sampling by skin diving?
Recently, I collected qualitative samples of marine soft-bottom zoobenthic communities at 10-15 meters depth by skin diving. I used hand-operated corers of 8cm diameter. I wonder whether there is any publication/report/manual on sampling procedures by skin diving.Following
Claire Wordley added an answer:Can we calculate Convex Hull volumes with categorical data?I am calculating Convex Hulls for high-dimensional trait data for a set of communities (Functional Richness) to understand how species pack and fill trait space. I was wondering if these calculations can be done using categorical trait data or are the analyses affected by not having a matrix of continuous values? Is there a way to circumvent this issue?
I also managed this using PCoA on Gower distances as suggested by http://villeger.sebastien.free.fr/ , it worked well.Following
Madhukar Baburao Deshmukh added an answer:Which are the best methods to remove termites?
We have been working with a termite species from the genus Odontotermes. We'd like to permanently remove termites from some mounds to observe their potential cascading ecosystem effects. Has anyone tried to do this?
It seems like one option to remove termites would be with insecticides, but we're worried this effect would only be temporary and termites may recolonize. One thing we've seen is people using plaster of Paris as a way to study mound architecture. Do you think we might be able to use plaster to permanently "plug" termite mounds? Anyone have any experience with these methods?
To remove termites you can use organophosphorus pesticide in the form of chloropyriphos spray or Mix. chloropyriphos formulation with soil or infected area.Following
Marcus Fritze added an answer:Can bat population reduction due to White-nose Disease or other factors be linked to changes in guano communities that rely on bats for nutrients?
Besides our concern over bat populations for their own distinctiveness and inherent value, we must recognize that bats support unique communities of cave life through their guano deposition. Declines in bat populations due to new stresses, including White-nose disease, would likely result in loss of biodiversity and/or abundance of life in cave environments. Is anyone looking into this directly? Can anyone suggest new study sites or approaches to test this in future research?
Paula Meli added an answer:I'm looking for a "plant trait similarity index", any suggestions?I would like to test how dissimilar are some traits of an intruduced species with respect to an established community.Following
Rocco Labadessa added an answer:How can I quantify the edge effect on plant communities?
Values of plant species cover have been sampled at an increasing distance from the edge of a grassland patch.
I was thinking about methods enabling to quantify the community change along such a gradient. What would you suggest?
In order to compare different edge types, is there any specific measure for the magnitude of the edge effect?
Have you ever addressed this issue or could you recommend publications in this field?
Dear, thank you all for your replies.
I wish to specially thank Audrey and Andy for their thorough considerations and useful suggestions...the discussion is getting even more intriguing.
This also leads me to hope for a cooperative research.
Nathalie Niquil added an answer:Is it possible to do a Fuzzy Correspondence Analysis (FCA) under constraint, like a CCA (Canonical Correspondance Analysis)?
Hello, we are working on biological traits analysis with fuzzy coding and would like to make the link with abiotic conditions.
How do you do it?
Thank you for your answers !Following
Mrigakhi Borah added an answer:How many days are required by Cyclosia sp. moth to complete its life cycle?Especially the caterpillar stage remains for how many days, can anybody tell me?
That means it depends on the climatic condition of the area.Following
Thiago Gonçalves-Souza added an answer:Does anybody know if it is always necessary to use a p adjustment for Fourth Corner Analysis?
In the exploratory analysis of my data, I see a clear pattern of relationships between certain environmental variables and species traits. When I carry out a fourth corner analysis it points out to those relationships, but with the p adjustments the significance is lost.
Nina Sajna added an answer:If higher stability entails higher diversity, how come there is low diversity in climax communities?Climax communities are said to be in a state of equilibrium because organisms have already adapted to their environment and succession is no longer taking place. Therefore, it can be assumed that it is stable. If climax communities have high levels of stability, why is it low in diversity?
there are also differences whether you look at plant diversity or if you look at animal diversity. Plant communities might have lower diversity in late successional phase (especially in the process of secondary succession), while the diversity of animals almost always increases (btw. I would be grateful for any reference contradicting the latter).Following
Giovanni Zurlini added an answer:What is the best diversity metric in community ecology--species richness, Simpson's index, Shannon's index, or another metric?There are many metrics for measuring biodiversity in community ecology. It seems most diversity metrics are based on the spatially nested hierarchical organization of regional biotas. I am wanting your opinions about what the best metric is for measuring community diversity and comparing such patterns across scales. There are probably many squibbles about this, and some people might for example say that Shannon's index is irrelevant and shouldn't be reported. What do you think?
Dear Francesco, but once you get the "True" diversity and decompose it into its alpha, beta and gamma components what are you going to do with it? I am really curious to know.Following
Philip Phil-Eze added an answer:Does anyone have experience using the Shannon Wiener Diversity Index Score?
I just completed the shannon Wiener diversity index and calculated a score of (H)=2.154890613. The values range from 0 to 5, with common ranges usually between 1.5 to 3.5. Can I really say that the population is diverse? What other index should I use when getting a score like this? Thanks
The diversity index is a dimesionless value. On the face of it, one can say whether diversity is high, i.e tending towards the upper limit of 5 or diversity is low, i.e. tending towards the lowest level of 0. However, it is important to relate diversity with the taxa under study, the abundance of population of individual species and the eveness of the distribution. Simple interpretation of Shannon Weiner diversity index as high or low is too elementary an analysis. refer to the references from Reza.Following
Stamatis Zogaris added an answer:What kind of microhabitat for birds could be in a riverine-estuarine-marine- wetland complex?In a coastal wetland with mangroves, lagoons, macrophytes, dunes, sea beaches, rivers overflow planes, etc.., You might consider each of these covers as habitat. Also in those habitats are presented gradients of salinity, oxygen, depth, currents, height of vegetation, sediment type, etc. Is there a classification of microhabitats for wetlands?
On the other hand, what kind of fine classification can I use for separate the birds (more than family, diet, etc ) to reflect the diversity of microhabitats in that wetland? In the wetland there are birds from different families, waders, swimmers, etc., many species recognized for their diet specialization, even within a trophic classification. For example, piscivorous have speciality in the size and the way to capture. Is there a classification that allows for a finer separation into groups like ducks (more than swimmers and divers), herons, waders, gulls? Or within groups as piscivorous, or filter feeding birds? References and publications links will be welcome.
Here, I attached a publication have I reviewed.
Thank you very much.
This is a great question. There are very simple ways to do this by building lists of species per habitat-type area. Again this is simplistic, but it may systematize the varied world.
If you have anything published please post.
Best of luck,
Atena Eslami added an answer:Does anyone know whether there is a site that had transparent pictures of salty glands in plants or not?I have been accumulating some papers for a seminar, but unfortunately I don't have enough pictures of that. Can anyone help me?
thanks for your answer but would you please send me the chapter which related to the salty glnds?Following
Graeme Ure added an answer:Does anyone knows how can we avoid pseudoreplication in a community ecology study?
Any minimum distance for each plot/ treatment?
If we take Travis’s example 3 lakes and 15 samples each. Actually each lake is a treatment. If the 15 samples are randomly distributed then you do have n=15. If however to get your 15 samples you decide that you can save time by going to 5 locations and take 3 samples at each location then those 3 samples are in effect sub-samples and need to be combined (as with Trine’s kelp) so n=5. By the same measure if you decide to get your 15 samples by taking 5 samples at the same place on 3 days then again n=5. If you decide for convenience sake to have 15 random samples but only on the side where there is road access, then you would need to be sure that the conditions on that side are representative of the lake as a whole.
For your animals the overlap question does get important particularly if your sampling methods are destructive (pit-fall etc). However, if they are not destructive and you want to either know presence/absence or trend then total independence may not be so important. A case would be ‘Five Minute Bird Counts” Bird territories (across species) are so variable as is behaviour that the most important thing is to be consistent with as many things that can be controlled as possible. You will double count some birds, equally on another day the more mobile birds may not be counted at all, not because they’re not there just because the 5 minute count didn’t coincide with their movements around a much bigger territory. Over time a population trend still emerges.Following
Ian C. Duggan added an answer:Does anyone know a good (user friendly + reasonable cost) ecological software for advanced community ecology analysis?
Currently I use PcORD Version 5. I am interested on PRIMER 6 now but keep looking for other options.
I use CANOCO or PRIMER, with the program dependent on the data. If you have gradients in assemblages responding to underlying gradients in environmental variables, then the analyses in CANOCO are best. If your underlying environmental variables are categorical (e.g., before/after, high/medium/low), then the analyses in PRIMER are best.Following
Alan M Kuzirian added an answer:Can anyone suggest histochemical changes during the reproduction in molluscs?I am working on the reproduction of molluscs and there are many articles which describe the histomorphological changes during the reproduction, but there are not any articles associated with the histochemical alterations in the body of molluscs during the reproduction. Can anyone suggest any articles on this?
In the early 1970s I described in extensively the reproductive histochemistry in several nudibranch molluscs. At the time, no one published master's theses, but the work is available and some of it was incorporated into other publications.Following
About Community Ecology
In ecology, a community is an assemblage of two or more populations of different species occupying the same geographical area.