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I have a pool of data in two different ecoregion (eco1 and eco2), each of one is a transect, 25 transect in one ecoregion and 25 in other ecoregion.
I want to compare the diversity of each one.
When i'm perform the analysis, i calculate the shannon index for each transect of all pool of data and later i filter the data for the 25 transects of eco1.
Is correct perform an analysis based in the average of shannon index of this 25 transects?
or the right form is calculate DIRECTLY the shannon index for the sum of all transects of one ecorregion ? and not based on of mean description
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Hazlo como recomienda Andrew Paul McKenzie Pegman , o usa rarefacciones. Eso se puede hacer usando el software PAST 4 or EstimateS (ambos gratis). Claro, el R tambien lo puede hacer.
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I am planning on bringing some plant material (leaves of carnivorous plants) from Ireland to the UK, the leaves will be stored in buffer/ silica.
One of the plants is CITES listed I know I will need the permit for that but I am unclear what if any phytosanitary/ other permits or certificates I will need to bring the leaves over. Most of what I read on government websites talks about bringing in live material or restricted plant parts (goods requiring prior notice).
If anyone with more experience with this can help me with a list of what I should make sure I have filled in that would be greatly appreciated.
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I would recommend that you contact the Horticultural and Plant Health Division of the Department of Agriculture, Food and the Marine (DAFM) (In Ireland) and Centre for International Trade (Foss House) Animal and Plant Health Agency (APHA), a branch of Department of Environment, Food and Rural Affairs (DEFRA) (In the United Kingdom). Each agency is involved in the process and certification of moving plant material and potential consequences of this. If you provide each agency (via their contacts pages) with information about what you intend to send they will provide you with the appropriate documentation and permits that you might need. You'd also need letters of recommendation or similar from the company/institution/department that you're sending from and is expected to receive it while doing the actual shipping process.
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Foresters usualy use a clinometer to calculate the tree height. These can be quite expensive to buy, but I don't know if using smartphone apps is accurate enough for measure canopy height. Do you recommends using automatic clinometer apps for field research ?
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I was using "Smart Measure version 1.7.7" for measuring height of trees planted along streets within settlements. I am moderately satisfied with the precision. I found the optimal distance to carry out the measurements from a distance about 2-4 times of the tree height. In most cases, however, I was not able to measure from the optimal distance because buildings, parked cars, etc. made it impossible.
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I recently began to work with viviparous Neotropical skinks of the genus Mabuya. Specifically with females at different stages of gestation. My tutor and lab colleagues have studied them for a long time and a recurring comment is related to the difficulty of field sampling.
The standard method of catch is by hand, but these lizards are very quick moving through the litter, and their smooth-scales covered skin makes them difficult to hold. Also, in the most advanced stages of pregnancy, these lizards stops feeding so funnel traps will likely be less effective.
I have little experience in catching, and I am planning some field trips to obtain some specimens (especially to learn about the field work). I would like to try different catch methods, hoping to make it easier to obtain research material.
I will be very grateful for any suggestions or advice you can provide.
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Artificial retreats :)
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For many camera trap studies it is a problem that it is not possible to distinguish individuals, therefore consecutive visits may be the same individual. Either an individual triggers a burst of captures during one visit, or the individual moves consecutively in an out of the camera view during a short time period.
To minimize double counting many studies are using a time gap (hit window), which is the length of time used to group consecutive images/videos together as single detections (given that the data are from the same camera and featuring the same species). This will vary between study designs, for example bait site will cause individuals to stay within the detections zone for a longer period compared to non-baited sites. Further, some species will be more likely to stay at the site than others.
Is there a good procedure to estimate an optimal time gap (hit window) for an existing data set?
Best regards,
Ronny
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This depends mostly on the methodology (baited/non-baited; random samples/non random) and especially the analysis method.
For most of the more recent density estimate methods very short (2s etc) or no time delay have been used.
If you have enough saving capacity and battery life (which has improved in newer models) I would say you should go for the shortest interval possible. If required you can remove/filter multiple triggers in analysis. Otherwise you may miss critical data during the time interval where the camera stays innactive. And I personally prefer video captures instead of snapshots (which helps better identification).
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We have set various in situ experiments with epiphytic orchids seeds. We put fresh orchid seeds inside nylon mesh packets (ca. 1000 seeds per packet) along with a bit of moss (to improve moisture), and then located those packets on tree branches close to mother plants. After 1 year, we retrieved the packets and open them to locate germinating seeds, but moss and lichens have grown inside of the packets, plus there is a large accumulation of detritus and dirt, so it has been very difficult to locate the seeds (only finding <5%). We don't expect mortality/decomposition rates to eliminate 95% of seeds.
Do you have a recommendation on how to locate those seeds?
We have tried the following:
1) series of washes and filters to remove bigger pieces of moss and lichens
2) washes and low centrifugation
3) centrifugation with filters
4) dilution of centrifuged materiales in several petri dishes.
We wish to use a method that wont damage the putative fungi growing in the germinating seeds / protocorms.
Thank you!
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Germination and seedling establishment in orchids: a ...
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I'd appreciate advice about acoustic recorders for undertaking surveys for a range of taxa, particularly birds, frogs and microbats (i.e. audible and ultrasonic). The units will be left in situ for days/weeks and will need to withstand a range of environmental conditions (e.g. deserts and wet tropics). I've previously used Song Meters. Is anything better.....that's not considerably more expensive? If not, which Songmeter model would be best? Thanks in advance.
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Hi, you can use the 4th generation Song Meter SM4 which is a compact, weatherproof, dual-channel acoustic recorder capable of capturing large amounts of data from wildlife such as birds, frogs and aquatic life.
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We are planning to sample crustaceans from intertidal area during low tides using transects. We want to catch all within the transects. However it's difficult to catch them because they can escape very fast. Is there a method to catch crustaceans easily? We have tried to use clove oil as an aenesthetic but it doesn't seem to be effective. Maybe there are other aenesthetics we can use. Any suggestions are welcome.
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Dear Al-Delemy and Mr. Kovalchuk
Thank you for the advices. So I should use various methods at the same time to be able to catch all the Crustaceans. 
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At times I catch 2000+ mosquitoes in a CO2 baited trap. I would like to get an idea of the population without identifying and counting every mosquito.  What sampling methods are appropriate? I have been quartering the pile and multiply the results by 4, but is there a better way? And is there a sample size (sample size formula) I can use to ensure I have a large enough sample to accurately represent the trap population?
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Thank you very much for your response.
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I found several R packages to do Bayesian GLM analysis. One of those is "arm". I like this package because it is simple to use. Nevertheless, I would like to see some articles where this package and function "bayesglm" were used in the field of ecology/forestry.
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Yes, R can be written to achieve any analysis. You could have a go at writing the code yourself. Then you can publish the code alongside your research! :-) 
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Hello,
I am a student in ecology, currently conducting a monitoring survey of butterflies in a reserve in Portugal.
I'm using a modification of the Pollard transect method in order to inventory butterfly species. My goal is to compare the butterfly diversity between the Inside and the outside of the reserve.
I have 3 different transects inside the reserve, and 3 transects outside, carefully defined to represent different types of habitat. I'm planning on doing 5 times each transect (so 5 replicates per transect).
At the end of my survey, I will calculate diversity indices (Shannon, Simpson,...) for each transect.
I was planning on doing a repeated measures anova to compare these indices, but I read recently that such indices needed to be transformed in order to be compared (for example, I should use exp(H') instead of H' in my statistical analysis).
My question is : do you think it's necessary to transform the diversity indices I'm going to calculate ?  If so, do you think a repeated measures anova is the good way to analyze my data, knowing that I'd like to compare butterfly diversity on different levels (Inside/outside of the reserve, but also between each transect) ?
I hope that my question is clear enough (if not, I can add some clarification).
Best regards,
Antoine
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Dear Antoine,
For comparing diversity Index (for e.g. Shannon) you can use the T-Hutchenson (Zar, 1996; page 157). It is a specific analysis for comparing diversity index, I attached to this message a .xls file for calculating Hutchenson tes. You have to do this in paired way. Ifyou need help I'am available without interest to help you. (by the way, you dont need to replicate measures, nad the anova it does not work for comparing diversity index).
Best regards
Carlos 
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I want to tag some gorgonians corals in order to return months later and see how much has it grown. I'd like to know if there's any specific way to do this. Is there a type of ink that doesn't damage the coral? I was thinking in making a knot with yarn, perhaps. 
Many thanks.
Regards
Bernabé Moreno
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I agree with Jérome Mallefet, better putting a tag to the hard substratum at the base, depending on the conditions it may be used a two component adhesive glue that works well underwater. I attach a sample of how the growth can be controlled with photographs, the measurement can be assured by putting a metric strip into the frame.
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I would like to use “Pollocks Robust Design” for a CMR study of endangered arboreal snails and would like some feedback on my approach. 
Collection surveys will be 2 sets of day-night-day data collection with 3 weeks in between events.    Or would “better” data be collected with more visits? The snails are  sedentary during the day- easier to spot at night using lights while they are moving. So, we know we find more snails at night. Is it ok to mix up the time of day that the data collection occurs?  
Survey Area is about ½ acre of enclosed habitat but the snails are concentrated in hotspots within this area. Should I survey those hotspots to get the maximum number of individuals or should I choose random quadrats within the ½ acre with the chance that the quad will have no snails. If quadrats then how many? I am planning to do a timed collection (one man hour for each area) but should I search to exhaustion in each section instead? One of my goals is to determine the approximate number of snails in this enclosed area but much of the habitat is too tall to be searched
I am planning to use Program MARK for the data analysis- any guidance regarding doing Robust Design analysis is welcome. I have worked my way through MARK before for closed capture CMR.
Thank you for any feedback.
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Just a few thoughts. If you want estimates of N, make sure your survey areas are consistent. If you want estimates to apply in a general sense to your area of interest, but cannot exhaustively survey the whole thing, then you need to employ a probabilistic sampling approach. You can account for detection heterogeneity in time of day by using that as a covariate in your analysis. Otherwise you will have unmodeled heterogeneity resulting in overdispersion. The more quadrats and sampling you do, and the better estimates of detection you get, the better your estimates of N etc. will be. Make sure to model average for N.
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I am currently working on evaluating the effects of climate change on marine plankton using a mesocosm facility. I have some queries
The capacity of my tank is 2000 L - Indoor facility
1. Do I need a wave maker for recirculation in the tank or is it enough if I use a high capacity blower (e.g. 80 HP).
2. What kind of instruments should I used for CO2 diffuser?
3. I am planning to run one set of experiment for 20 days, the total capacity of seawater will be 2000 L, Do I need a separate recirculating system?
4. I will be drawing around 10 L for various analysis on alternate days for a period of 20 days.. how much will this effect the total volume ? I don't think refilling is a good idea, kindly suggest me some alternative.
5. What are the general problems that we could face while using a mesocosm facility?
Thank you
Regards
Vinitha
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Hello Vinitha, 
If i may suggest some literature that you will find very useful and will answer some of your questions:
The Guide to Best Practice in Ocean Acidification Research:
A methods review from Dr. Cornwall et al., 
1) You do not really need a wave maker, but a recirculation pump, unless of course wave motion is a parameter that you want to add to your experiment;
2) For CO2 diffusion it is advised to use an atomizer, but strongly depends on what kind of tank (e.g. if your tank is closed or open) and on your CO2 injection systems. Also in your case with a 2000L tank you will need A LOT of CO2 to maintain high CO2 to simulate future oceans conditions, that will be hard (but not impossible) and costly to accomplish;
4) if you have a closed system, you have to consider refilling... using UV filtered seawater, or a synthetic one that you can DIY
5) there are many problems that you face, and these are similar to the ones that aquarist and aquaculture professionals experience, so you will find a lot about proper methods of cultivating planktonic communities. For example, water changes, proper nutrients concentration, oxygen depletion, in your case also CO2 diffusion, temperature maintenance, proliferation of unwanted organisms... etc etc... 
personal advices:
I just wonder why you need so much volume, since you need to replicate your experiment, there are simpler (and easier treated) tanks that are designed for planktonic communities, for example a very very simple one that you can build yourself: http://www.reefkeeping.com/issues/2002-07/ds/images/image002.jpg 
Hope it helped you a bit, this is as far as i can go with the information you gave, 
Regards, 
Fabio
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Leaf Area Index (LAI): was calculated as ratio of the leaf surface area to the ground area occupied by a plant stand (Thomas et al., 2003). = Leaf area / ground area.
Through this way you can calculate the land area of selected  crop plants of your treatments. Same calculation will be measured at 30 and 60 days in selected plants or according to your trial,   
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I know that there are methodologies different, but I would like to know a reliable mode for this specie and for calculating the % volume of prey inside it.
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Tahnk you so much at all
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Does anybody know how many underwater photo quadrats (1mx1m) are needed for a 20m transect line?
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The answer is not that simple... We could start by why 1 m x 1 m quadrats and why a 20 m transect line? Even that could be discussed. And a transect over what? How homogeneous or heterogeneous is what you want to sample? Power and sample size? Statistics you will use?
I recommend that you read:
Chopin, T., 1997 - Marine biodiversity monitoring. Protocol for monitoring of seaweeds.  Environment Canada, Ecological Monitoring and Assessment Network, Ottawa, 40 p. 
The text is available on my ResearchGate page.
All the best,
Thierry Chopin
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i want to design a temporary monitor of elephant movement between nearby communities, how can i mark a group without capture and make every resident recognise the group, spraying paint by remote control helicopter seems expensive for me.
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I agree with Salindra. I understand that ear characteristics, such as tears, ear venation, and tusk size and shape have been used to identify individual elephants (see Whitehouse & Kerley 2002. Oryx 36, 243–248, and also the associated PhD thesis). I would be doubtful whether paint would be useful for a long term study. Elephants will scratch and rub themselves against trees, alongside wallowing and washing behaviours, I can't imagine the paint lasting long. Also, if a semi-permanent paint is found, there are some ethical questions about whether we should be marking animals with unnatural colours for long periods.
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I'm planning an undergraduate student field project with mark recapture on wild mice and voles, and thinking about marking methods. As individuals only need to be marked for a week I don't want to use tags. I know fur clipping is often used, but what about temporary marks with hair dye or food dye? As this is for a student project with minimal previous experience handling small mammals I'm a bit cautious about using clipping. If you have used food dye / hair dye or know of something published which did, which brand was used (or a link to the reference)?
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I used nail varnish to paint the toe nails of small mammals. I got the best results when using fast drying ones with glitter inside.
But if you want to recognise the animals after more than 3 days I would prefer fur clipping!
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I just want to know about any marking technique for water monitors to identify an individual from a distance.
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Whatever you do don't paint their heads! Attaching anything to the animal is going to snag, drop off or hinder growth eventually. Recognising pattern on monitor lizards is very tricky because they are often covered in dirt, shedding skin, in the shade etc. It looks easy when they are wet and freshly shed but it's not a practical way of doing it. By far the most efficient way I have found for biawaks is to cut a unique set of notches into the tail crest with a very sharp sterile blade. If it's done carefully they can be made deep enough to be permanent without hurting the animal (you can tell very easily when a biawak is hurt by its reaction). Never cut beyond the crest! These pictures show tail notches made 1-3 years previously. Mid third of the tail is best because the crest is high and they are very unlikely to lose that much tail. Sorry for late reply.
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I am writing a review of hunting sustainability assessment in New Guinea (Indonesia & Papua New Guinea) and would like to find out if there are any studies conducted in Indonesia that have used actual sustainability parameters/indicators to assess hunting sustainability and not just speculate on hunting sustainability.
There are tonnes of articles on hunting sustainability assessments in the Neotropics and Afrotropical countries that have used sustainability parameters/indicators to assess hunting sustainability with a few in Papua New Guinea, but I can't seem to find any in Indonesia. Hunting sustainability assessments conducted in Indonesia are predominantly based on hunting yields, trade and other variables  to speculate or make inferences to hunting sustainability
I would like to find out if there are any unpublished work out there that have used actual sustainability parameters/indicators to assess hunting sustainability in Indonesia.  
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I have this paper, however it is only a speculation on the sustainability of the commercial trade of pythons and not a direct measure of sustainability of wildlife harvest nor did it use any or some kind of harvest or production model to assess sustainability of wildlife harvests. I am interested in studies that have actually used harvest or production models to assess sustainability of wildlife harvest (hunting).
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I'm capturing video of mosquitoes feeding on nestling birds in an attempt to quantify biting pressure(sample video is attached), but I have no way of determining which species of mosquitoes are attempting to feed.
What kind of equipment do I need in the field and what kind of software is needed in the lab to determine the species present?
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You would need a sensitive microphone very close to the chicks but I don't believe you could identify species from the sound alone. Much easier to identify from the videos. If they are Culex, pipiens had a higher frequency than tarsalis when we recorded them in the lab.   Peter
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The Bushnell Natureview ND Max camera with IR night vision might do the trick, but does anyone have any experience with recording insects in challenging field situations? 
I do not have a huge budget and the risk of equipment loss in my study context is quite high. I was wondering if there are any cameras out there designed for other purposes that could be adapted to remote (battery operated) macro-recording of night and daytime insect activity?
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Hi Peter,
I would have a look at Favaro et al. 2012. TrapCam: an inexpensive camera system for studying deep‐water animals. Methods in Ecology and Evolution 3:39-46. While it might seem a stretch, Favaro designed this himself to look at prawn catches (which you could equate to insect-sized animals). It was cheap and dirty, but it worked really well and the footage he got was excellent.
Hope this helps a bit.
Aleks
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Hello everyone!
I have a question from our mammologist group: they counted the number of anthills destroyed by bears on 89 transects (800 m each). They had a huge SD: 23.06, and the mean of destroyed anthills per transects was 27.4.
How should we calculate the minimal number of transect needed to monitor the bear activity reliably?
Thanks,
Attila
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ISSUE: Minimum sample size
MINIMUM SAMPLE SIZE TO CONDUCT STUDY: In general, if the population size is known, the Yamane method is used to determine sample size. However, the method is not efficient, i.e. it tends to requires larger than necessary sample size and it cannot handle unknown population size scenario. See below for citation.
Assume that the total population (N) is non-finite, if we need the minimum sample size to study the ants in this case the following formula may be used:
nmin = (Z2σ2) / E2             ... where E = σ / sqrt(nobs)
... where Z = 1.65 for 95% confidence interval; σ = 22.92 using the t-equation to solve for the estimated mean (μ = 23.39) and the Z-equation to solve for σ, thus:
t = (X^ - μ) / (S / sqrt(n))
... where X^ =27.40; S = 23.06; and t = 1.64 using 0.95 CI for infinity degree of freedom. Solve for μ. The solution is μ = 23.39.
Z = (X^ - μ) / (σ / sqrt(n))
Substitute the value of μ into the Z-equation and solve for σ. The solution is σ = 22.92. Now put the values into equation (1) and solve for nmin. The solution is nmin = 242.18. This is the minimum sample needed to conduct a study of this population. The confidence interval used was 95%. If you need higher confidence interval, change the relevant value for t and Z according to the confidence interval level.
REFERENCES:
[1] Yamane, Taro (1967). Statistics: Introduction to Analysis. New York: Harper and Row. p. 886.
[2] Montgomery, C., Runger, G. C. and Hubele, N.F. (2001). Engineering Statistics, 2nd ed. John-Wiley, ISBN 0-471-38879-3. p. 172.
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As specified in national and European laws and directives (INSPIRE, Aarhus convention…), ecological data in Europe must be accessible and free for use by the research community as well as other stakeholders. Scientific research questions in ecology can be resolved at local, regional and global response scales by concomitant stakeholders only by combining data of different disciplines. Do you know any review or list of existing databases in your own disciplinary field in ecology?
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Given your focus on ecology, you may want to start with this GEO BON document:
The GEO Secretariat (https://www.earthobservations.org/index.php) is dedicated to the coordination of data acquisition and analysis in a wide range of thematic fields called "Societal Benefit Areas (SBA)", and strongly advocates open access and inter-operability.
The weather and climate community has a very long history (>150 years) of developing common measurement protocols and sharing data, so you may gain insight by investigating the work and findings of
- the Global Climate Observing System (GCOS: http://www.wmo.int/pages/prog/gcos/), which coordinates and oversees efforts to acquire, archive and distribute climate data worldwide, in particular for atmospheric, oceanic and terrestrial applications. Look, in particular, at the various pages under "About GCOS", "Observing Systems and Data", and "Outreach" (in this latter case for a list of publications).
- the World Meteorological Organization (WMO: https://www.wmo.int/pages/index_en.html) and specifically the Global Framework for Climate Services (GFCS: https://www.wmo.int/pages/governance/ec/global-framework-for-climate-services_en.html).
Another large, multidisciplinary organization to monitor is the International Council for Science (ICSU: http://www.icsu.org/), including its flagship programme called "Future Earth" (http://www.futureearth.org/).
All these international structures share a common interest in database structure, inter-operability, effective and open access, etc., not only to stimulate interdisciplinary research but also to promote the sustainable development of nations and societies.
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I've just read about this concept, but I can't find any exact definitions for it. I'm not working in the field of ecology, so could you please help explain this term in brief?
Thank you so much.
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Hi Danny, delayed density dependence refers to density-dependent mortality that involves a time lag after the increase in host/prey density and subsequent predator-induced mortality. For example, in souther pine beetles, the populations increases rapidly, but it takes a year for their predators to produce a new generation, so there are cyclical population trends where beetle abundance increases dramatically and then crashes a year later when the predator population catches up. This is also seen in some fish, where small, fast-growing species rapidly reproduce and then crash a year or more later, when larger, slower-growing predators finally start to limit the population of small fish.
For a better explanation, check out Hajek, A.E. 2004.  Natural Enemies: An Introduction to Biological Control. Cambridge University Press.
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I am looking for a GPS unit helpful in field biology. A handheld unit, affordable and easy to carry.
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I usually work with Garmin 60Csx and 62Csx, like Galo. But now there is a new option Garmin GPSmap 64st . All of them are relativily cheap, weight is ok, and the new one has a very good improved signal (GPS+GLONASS). 
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Asociacion Armonia is protecting a 11,000 acre tropical savanna in the Beni, Bolivia. We are going to use a savanna patch-burn management technique, but we are not clear of the potential climax level of older successional savannas. We are thinking the best measurement could be arthropod diversity and abundance. We are interested in an comparison arthropod studies in tropical savannas.
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Probably some of these: 
Frizzo, Tiago L. M. ; Campos, Ricardo I. ; Vasconcelos, Heraldo L. . Contrasting Effects of Fire on Arboreal and Ground-Dwelling Ant Communities of a Neotropical Savanna. Biotropica (Lawrence, KS), v. 44, p. 254-261, 2012.
FERREIRA, R.N.C ; Franklin, E. ; Souza, J. L. P; Moraes, J.. Soil oribatid mite (Acari: Oribatida) diversity and composition in semi-deciduous forest fragments in eastern Amazonia and comparison with the surrounding savanna matrix. Journal of Natural History, v. 46, p. 2131-2144, 2012.
SILVA, LAURA V. B. ; Vasconcelos, Heraldo L. . Plant palatability to leaf-cutter ants (Atta laevigata) and litter decomposability in a Neotropical woodland savanna. Austral Ecology (Print), v. 36, p. 504-510, 2011.
Lopes, Cauê T. ; Vasconcelos, Heraldo L. . Fire Increases Insect Herbivory in a Neotropical Savanna. Biotropica (Lawrence, KS), v. 43, p. 612-618, 2011.
AMPOS, Ricardo Ildefonso de ; VASCONCELOS, H. L. ; ANDERSEN, A. ; Frizzo, Tiago L. M. ; SPENA, KELLY C. . Multi-scale ant diversity in savanna woodlands: an intercontinental comparison. Austral Ecology (Print), v. 36, p. 983-992, 2011.
SANTOS, E. M. R. ; FRANKLIN, E. ; MAGNUSSON, W. . Cost-efficiency of Sub-sampling Protocols to Evaluate Oribatid Mites Communities in an Amazonian Savanna. Biotropica (Lawrence, KS), v. 40(6), p. 728-735, 2008.
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I’m working on a paper considering the role of facilitation in niche theory and would appreciate comments, suggestions and criticism of the following. In theory the niche is operationally defined abstractly as an n-dimensional hypervolume that is impractical if not impossible to really measure. In practice, however, in field studies and extinct community reconstructions the niche is typically operationally defined and measured as habitat space occupied with and w/o neighbors (fundemental and realized niches respectively). How does one cope with this disparity between theory and reality? Are field and fossil studies that equate habitat distribution with niches bogus?  Is the rigorous n-dimensional definition of the niche a theoretical construct that is virtually impossible to quantify in complex communities? or Does the answer lie somewhere in between?
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To stimulate more discussion:
The relative concept of the 'niche'
Some people claim that different species do not share the same 'niche', for instance defined as an X-dimensional or endless-dimensional space where individual species live. Some would even claim that there is no overlap in the niche across species because of natural forces such as inter-specific competition. However, the level of niche overlap between individuals or species will probably depend on scales of analysis. For instance, unicellular or multi-cellular individuals or species should express niche overlap when the niche is defined with a small number of variables. Each unicellular or multi-cellular individual probably have their unique niche when the niche is defined with X-dimensions or endless dimensions.
Is the size of the niche perception-dependent or place-dependent?
The definition of the niche will evidently depend on what variables are empirically studied. If only one dimension is used to define a niche, such as the place where an individual organism lives and moves, does an individual vertebrate and an endoparasite (e.g. ancient mitochondria) living inside an individual vertebrate share exactly the same niche or not? Both the vertebrate and the organism living permanently in or on the vertebrate move together and therefore may share the same niche. However, the average physiological environment of the endoparasite will not be the same than the average physiological environment of the vertebrate, so individual endoparasites and individual vertebrates would not share the same niche from a physiological point of view.
               
If information mentally perceived are defined as resources exploited by living beings, does this imply that the size of the individual niche depends on the spatiotemporal scale of perception mechanisms involved? The perception niche of physical contact will probably be substantially smaller than the perception niche of a visual system or the perception niche in dream expressions. How to identify dynamic perception niches in wildlife, accepting scientists can never obtain detailed information provided by wildlife not been able to explain what they see, what they smell, what they hear, what they dream, etc.?
               
Different species should express significant overlap in niches when a given species explores niches of other species, for instance to find energy or material for living (soil, wood, plants, animals). If humans explore any environment in the world, for instance because of curiosity or to get resources for living, what fraction of the so-called human niche is shared with niches from so-called non-human organisms or species? Does this imply that both the individual-specific or species-specific niches are probably difficult to define with precision from an empirical point of view?
Could a niche be defined as a probabilistic place of living or a probabilistic space of perception occupied by an organism with a probabilistic design? Accepting that Earth as a niche of living is permanently moving either around its axis, is permanently moving around the sun, and is following moving trajectories in the universe, would this imply that the definition of the niche changes with research domain (e.g. astronomy versus earth ecology), also to provide additional arguments that individual niches are unique in time and place for at least one scale of analysis?
Accepting nothing is fixed an individual will always express continuous spatiotemporal variation in physical design for at least one level of analysis, and therefore could be defined as a probabilistic unit.
Niche differentiation across species exploiting artificial winter feeders
Artificial food such as fat balls hanging from a wire attract small passerines like great or blue tits or chickadees, but also attract other species such as robins or house sparrows having niches different from that of tits/chickadees. When a fat ball is attached on a wire 10-20cm above the ground, tits will use the fat ball as a perch to get food of which some is falling on the ground. Sparrows or robins will never touch the fat ball, but just wait under it until the food extracted by tits is falling on the ground. Why robins or house sparrows did not evolve simple niche shifts to exploit the artificial fat balls? Why is species-specific plasticity in habitat exploitation reduced, even for very simple food searching tasks, like extracting food from fat balls hanging from a wire? Do some species through their foraging activities provide food to other species, as illustrated with the case described above?
What is the size of a 3-dimensional garden, property, or territory?
Properties, gardens, or so-called territories are most often defined using 2-dimensional measurements expressed in length and width, but how to define a property, a garden, or a territory using 3 dimensional measurements? For instance, is the 3-D of a garden limited by the height of the tallest trees in the garden and the deepest roots in the garden, or not? Does this imply that 3-D of a garden continuously changes? And what if someone can build a ladder of more than 1 km high? Are the people that are climbing the ladder still in the same garden? What is the highest acceptable height defining the 3 dimensions of a garden? If an airplane is flying over a garden, is it flying in the garden defined with 3-D, or not?
Other questions
+ What are consequences for estimates of intraspecific population densities or interspecific densities influencing life-history expressions within species (e.g. lay-date, clutch size)? For instance, tropical forest research indicates that different species are observed at different heights in the same trees, which implies that different species occupying these trees may have the same territory or niche size at a two-dimensional level, but not at a three-dimensional level.
+ How to define a territory or niche when every individual movement is unique in time and space?
+ How far should we go in the past to define a wildlife territory of an individual? If history is not important in the definition of a functional territory, can the functional wildlife territory only be defined through current position and perhaps probabilistic future movements?
+ In an ever dynamic world, the individual functional territory probably equals the individual space of movement best represented by a network of traces on a map.
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I am wondering what solutions have you found to have your GIS data on the field with you? I do not mean simle points/coordinates, but also raster files (derived from remote sensing). I am interested in both hardware and software solutions. At present I use an Android tablet with Oruxmaps and Qgis for android. But they lack many features and are very limited
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Geopaparazzi for Android...and geopapazazzi plugin for QGIS....
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Anyone know how to keep samples at constant -20°C on the field for a couple of weeks?
I will not have access to dry ice or LN2, so I'm looking for a compact freezer.
The littlest I've found is 11 lt., but I will need just 0.5 lt (10-15 capillary tubes).
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You may use the archaic method with a mixture of solid salt in ice and saturated saline water. Eutectic point is reached at -23°C. This was used in the 18th century for the making of ice creams during spring/summer time out of saved snow. This should save you a freezer and energy problems.
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I have the Askew: Dragonflies of Europe, 2004. But the damselfly, Coenagrion ornatum, that I want to study is not in it. Can someone help me with a few good identification keys?
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Hi Anna,
- this book is probably the best guide for exuviae (and larvae) on the market (and includes the discrimination of Coenagrion ornatum from other species of the genus). You'll have to buy that, it's not available as a PDF, but it's worth the price. A bit of a problem is the Dutch language, but the numerous photographs help a lot to identify a species. There is also a very good guide by Steve Cham for the UK, however with less species included (and without Coenagrion ornatum). Note that the Dutch authors of the first book I recommend are currently working on an according guide for all European species, then in English language - we are all eagerly waiting for this publication.
Regards
Florian
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Will fixed camera sites be effective? If so, which model and make are economical and efficient?
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Thank you Kurt Rinehart
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I have field samples that I freeze in liquid nitrogen to stop all processes at a certain time point and want to use for transcriptomics. However, I will have to ship them which probably can't be done frozen since I don't know how long they will sit at the border so they might perish. Is it a good idea to keep them in ethanol for instance?
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we use dry shippers for that purpose. They hold -140C for about 21 days and can go on airplanes. They are expensive to get but once you have them they pay off