Questions related to Field Ecology
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
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.
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 ?
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.
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?
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.
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.
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.
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?
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).
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.
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.
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?
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.
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.
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)?
I just want to know about any marking technique for water monitors to identify an individual from a distance.
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.
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?
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?
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?
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?
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.
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.
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?
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
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).
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?
Will fixed camera sites be effective? If so, which model and make are economical and efficient?
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?