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How do i calculate the infiltration rate, Darcy velocity and pore water velocity for a field experiment, supposing i poured about 500 litres of my sample on a 3m by 3m field area of soil. I also took soil samples at various depths from 0-150 cm and had porosity for the different depths. So i need proper calculation and paper referenced on the method used.
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Thanks for your contribution
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This experiment would be conducted with:
1. Five paddy varieties
2. 4 levels of bio fertilizer (ex: control, 100, 200, 300)
3. A field experiment was conducted to study the growth and yield of rice.
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What would be the most suitable experimental design to select a most suitable type of bio fertilizer level for five paddy varieties? Based on the explanatory text of the question, my first suggestion would be a 5x4 factorial experiment in a randomized complete block design.
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i have read many peer review articles mostly they have written giving 200mm of water for X-crop in his total growth period. my question is that
200mm means amount of water in one acre or one hector or per plant in open field experiment?
please guide me about this matter
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Rubina Ansari thanks for your reply.
my question 200 mm is the amount of water giving per acre per plant etc because they are not describe it in details.
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Why did bio-aerosols not become a commercial alternative to silver iodide in cloud seeding?
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Going to investigate the use of SNOMAX for cloud seeding here in the West USA.
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It is the agri entomology experiment conducted in two different contrasting locations for two years and two seasons per year. The data collected from the experiment was number of eggs, larvae, pupa, adult insect
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I would suggests you can use multivariate analysis, correlation analysis , anova using sas software accordingly you can see the intraction effect.
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Which statistical test should e follow to significant difference between the treatments in field experiment, DMRT to Tukey HSD
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Thank you
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In a field experiment if there is possibility for both designs factorial RBD and Split plot design, and both qualifies as per degrees of freedom, then what design should be choose and why?
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The main difference between Randomized Block Design (RBD) and Split Plot Design is that, in the case of RBD, our purpose is to study the effect of one factor, which has different levels of equation precision for all levels. Suppose you are going to study the effect of different levels of nitrogen alone on the grain yield of one crop. In this situation, only the rate of nitrogen will vary, and the rest of the practices will remain the same for all the treatments. All treatments will be replicated an equal number of times for each treatment. In such a situation, the RBD design is recommended.
But what happens in the case of a split-plot? You want to study the effect of two factors, one with more precision and one with less precision.
The factor which is very important to you will be kept as a subplot, and the factor which is less important to you will be kept as the main plot.
For example, you want to know the effect of irrigation (irrigated and rainfall) and different levels of zinc such as 20 kg, 40 kg, 60 kg, and 80 kg/ha) on wheat yield. However, in this situation, your focus is more on zinc than irrigation, and you want to study it precisely so you will keep zinc as a subplot that will be replicated more times, so the result will be more accurate. Likewise, you will keep irrigation and rainfall as the main plot, and its effects will be studied with less precision.
Your treatments will be like this:
A: Irrigation (Main plot)
a-Irrigated
b-Rainfall
B. (Zinc) Subplot
20 kg/ha.
b. 40 kg /ha
c. 60 kg/ha
d. 80 kg/ha
I hope this has clarified things for you.
regards
nesar
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Drought treatments in field experiment include 1/12P, /1/4P, 1/2P, 3/4P , P. P represents the plot receiving natural precipitaion. I do a linear regression between drought treatments and response variables. The reviewer think that this analysis was inappropirate. Can you help me? explain why
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It makes sense to determine the trend over those treatments.
My opinion is that it would be fine to treat those treatments as quantitative treatments. Having five levels isn't ideal for this application, but I think it makes sense.
Another approach, though, is to conduct the analysis with those treatments treated as categorical treatments (e.g. as in anova). You can then apply polynomial contrasts to determine if there is a linear or quadratic trend over those treatments. This is a traditional technique in agronomic studies. The problem with this in this application is that the treatments are decidedly and intentionally not equally spaced.
This might be helpful with polynomial contrasts: https://www.ndsu.edu/faculty/horsley/Polycnst.pdf
Another idea: You might take a look at my paper at the following link. Here, we treated the treatments as categorical treatments throughout the paper, but then used them as the independent variable in a linear regression (Fig. 6). This approach might be okay with your reviewers.
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I am planning field experiment and wish to collect RGB as well as multi spectral images for crop monitoring.
Considering images shall be acquired by UAVs, I wish to know if any guidelines exists for conducting field experiment.
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There is no link on agronomic field experimental design with UAV based image processing. You can use RBD, which is commonly used for field experiment
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Hello statisticians in the house,
I am collecting samples from an already established field experiment with the following research details
Plant cultivars = Whole plot, 3 levels (cultivars A, B, and C)
Grazing level = Subplot, 2 levels (Grazing vs no grazing)
Block= 4 blocks
However, for my current experiment, I am only interested in the plant cultivars without the grazing effect. So, I collected the plant samples at no grazing level. How do I go about my statistical analysis? Should I treat it as a one-way ANOVA with blocking as a factor considering that there is no subplot factor anymore?
Thanks,
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Salvatore S. Mangiafico , I found a solution to this question. Since, the subplot treatments have been removed, the design is reduced to randomized complete block design.
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Many papers suggest having a minimum error degree of freedom of 12. When a new crop variety is evaluated for agronomic performance with a local check or control, number of replications needed to generate a degree of freedom value of 12 is 13. That is, number of treatments (n-1) X number of replications (r-1), which is (2-1)X(13-1). I maintain this minimum degree of freedom for experiments with three or more treatment, however I not sure in the case of a paired field experiment. I would be grateful for your support and suggestions.
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Sorry, I doubt that there is a standard sample size for this, I suspect it would just be bad statistics in any case. That might make some government agency happy, but would be meaningless. If you want to have enough power to detect a difference in yield, you must define that difference for a paired t test. Then you need a variance (or standard deviation) for that crop in your immediate area, climate and soil. Then, you will need to specify alpha and betta.
However, with a tiny loss in power, you could use Wilcoxon's paired sample test. Then you would only need to specify the minimum reasonable difference you seek as well as alpha and beta. I will leave looking up the calculation details to you,
My book has the details,
Research Decisions and Estimation With Confidence and Power: Up To Date Text and Reference From Classical and Recent Literature. ISBN-13: 978-1532721076, LCCN 2021913804
but my primary source for the sample size was:
Noether, Gottfried E. 1987. Sample size determination for some common nonparametric tests. J. of the Am. Statistical Association. 82:398. Theory and Methods.
PS: I haven't spoken Spanish since I was very young and I am now very old, But, Mamerto Reyes Hernández seems to be providing a similar answer and warning.
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Dear colleagues, I would like to ask you for advice. I would like to see if long-term field trials can answer the question of how a changing climate affects wheat yields and whether a changing climate will affect different localities in different ways. I have the results of yields from a long-term field experiment (1980 - 2018) and information about the weather (temperature and precipitation). However, a single field trial cannot provide such an answer. This is due to the interaction between the weather and the different varieties used in the experiment. It is therefore not possible to distinguish between the benefits of varieties and changing weather. But I also have the results from other experiments. We run three long-term experiments, each in different soil-climatic conditions. These experiments have the same methodology, the same fertilization treatments, the same preceding crops, and the same varieties of wheat used during the time. Would this be the way? On the other hand, each wheat variety will have different results depending on soil-climatic conditions. An opponent easily drops such an article from the table. The more I think about it, the more desperate I am. I wanted to somehow use long-term results, a long time series, obtained under defined conditions (experimental data), but the interaction between the varieties and the weather forms an impenetrable barrier. Can you think of an analytical method that would evaluate such data in a reasonable way? For your info, I tried to use different models and the cubic model provided nice results, showing a kind of stagnation (Ivanovice, chernozem), decrease (Caslav, Luvisol), and increase of yields (Lukavec, cambisol) in the last few years when warm seasons began. But again, different wheat varieties were used between 2011/2014 and 2015/2018. I have also evaluated if the season was cold, normal, or warm or dry, normal and wet. Thank you if you have read it and you have a tip on how to use and evaluate such data and provide the answer to the question of how climate change affects wheat yields.
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Dear colleagues, thank you very much for your tips!!! I will think about it and apply your advice!
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I'm looking for an algorithm to help automatically track soccer ball in a controlled field experiment (cameras positioned near to the ball) and output its time-series 2-dimensional coordinates. While we have found very efficient methods to do this in human movement features (e.g. OpenPose), we are experiencing some difficulties to found similar methods to detect/track other objects (e.g. ball). If you have some positive experiences with a given method, please let me known. Thanks in advance. Luiz H Palucci Vieira.
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Hello! Try to use Kinovea (free soft). Nice program for automatically tracking.
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The conservation agricultural field experiment with three treatments is being carried out in black cotton soil of central India for the last three years. Which statistical design is best suited for comparing two/three years of field crops and nutrient data? Treatment includes 1. Irrigation methods (2), 2. Sowing machines (3) and 3. Residue retention (3).
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@Himanshu Verma,@Subhash Babu, In these experiments, under main plot there irrigation two methods are there, under sub plot three residue retention levels and under sub-sub plot three sowing/planting machines are there. Experiments are conducted for three cropping systems viz., Rice-wheat, maize-chickpea and maize-wheat on one hectare of land each. All experiments are having three replications.
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I have a question regarding manipulation checks in field experiments. Can you please share if you have ever done one? Are you aware of any published studies that included one?
Usually, the price to pay for the natural setting of a field experiment is that we give up some control, including (I thought) manipulation checks. But lack of a manipulation check in a field experiment came up recently in a manuscript review and I would love to read and learn more about it. Input from all disciplines (especially social sciences) is welcome! Thanks!
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Interesting question. Following the discussion.
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Suggest me some related references.
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I agree with what Paul Milham has said. But ideally, in a field experiment with treatments laid out following a correct statistical design with 3 or more replications, a treatment should be tested at least for 3 years. If out of three years, results differ in one of the three years, it should be tested for at least one more years.
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I have some interesting data testing Kefir and some food effects in mice models of obesity and malnutrition. However, I just found Open Access journals and the Brazilian government it is not paying for that. Could you suggest me good journals without publications fee?
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Check LWT-Food Science and Technology
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Kindly suggest which statistical experimental design is suitable for a field experiment having two levels of irrigations (sprinkler and furrow), three levels of sowing machines (zero-till-drill, raised bed shaper-cum-planter and conventional seed-cum-fertilizer drill) and three levels of residue loads ( 30%, 60% and 100%) with three replications?
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From the context of the question it seems that you have three factors (irrigation, sowing machine and residue load). A first approach could be a three-factor completely randomized design.
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I want to analyse my experimental data of field experiment in botany,which includes plant growth.I want to apply standard error and critical difference? I want to know what all parameters do I need to findout the critical difference? And what is exactly Critical difference?
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It is the least difference greater than which all the differences are significant, also known as least significant difference.
It is used to compare the observed differences among different treatments. If the difference is greater than critical difference, it is considered as significant and vice versa
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Two questions:
(1) A person who constantly works on secondary data (in scientific research) is considered a scientist or data analyst?
(2) In a world of limited funding and "publish or perish", should scientists shift towards the secondary data approach?
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See the attached paper. It may help you decide the answer to your question. Best, David Booth
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I was studying plot-based comparison of two treatments regarding their contribution to sediment response. the slope is also another factor going to consider in the field experiment. the plot is designed in such a way that the runoff and sediment from the plots direct into the runoff collection tank. Here I want to take A sample of sediment concentration from the storage tank after each rainfall events occurred. but I am really confusing how to sampling and what equipment should I use while sampling. and also to what or at what depth should I take the sampling from the rectangular tank?
I want to really thank you for your support, guidance and your constructive suggestion regarding the issue.
Thank you!
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In my researchgate, there is a paper on Hazel Pistol Erosion Plots. I used 55 gallon drum to collect stormwater. As the water came into the drum, there was a container to collect what we called "heavies", like sand, gravel. That was measured separately from the drum contents. The level in the calibrated drum was measured for volume. The contents in the drum was water, silt and clay. A piece of plywood was cut slightly smaller than drum dismeter, about 2-3 inch holes were cut in the plywood, and attached something like a broom handle to the plywood, with added support from wires for stability. This was used to vigorously mix the drum contents, theoretically at least to a uniform concentration, then quickly grabbed a sample. The volume of the water in the drum times the concentration of the filtered sediment in the sample was used to get the "fines" contributed from that storm or sampling interval.
More recent was the study of sediment from a small, ephemeral gully paper. You might be able to use the geotechnical woven filter cloth to make some large filter collectors, which might collect and separate sediment from water, and possibly better than collecting just the quick to settle sand and gravel particles. If the filter fabric collector was preweighed, it could be taken back to lab, dried and weighed. The water in the tank could be stirred to check if there were clay materials passing through through fine filtration or the calibrated density float (forgot name) that can be used to measure the water plus sediment density, and that can be used to estimate the concentration amount of clay in suspension to be applied to the filtered water.
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Say you have 50 randomly selected flag leaf samples from each of your trial plots in a field experiment. You have their dry weights and you determined micro-nutrient and macro-nutrient content by ICP-AES. Is it possible to determine nutrient uptake efficiency, specifically that of phosphorus, from this data? If not, what type of information can we infer from this data?
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I know the height of the plant at different stages can be used to infer the mass. Regardless making this calculations would be interesting. For the best results a pre planned experiment with the pre set methods and purpose would be preferred. I still would make the calculations to see if something stands out related to the values.
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We have conducted a field experiment to evaluate the effects of infiltration trenches and improved grass species on biomass productivity and soil quality. The result indicates that water conservation through infiltration increased biomass productivity while the available form of nitrogen (ammonia and nitrate) content decreased, but the total N content increased. What would be the possible explanation? The reduction in available N could be related to the increased uptake by the grass, and possible denitrification due to increased soil moisture content because of infiltration trench. The organic matter content has increased with the increased biomass- can this alone explain the increase in total N?
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The increase in organic matter content enhanced the nitrogen availability and this is might be due to additional of plant origin material.
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Thinking in terms of a social setting such as a dance, a concert, a meal, if an experiment were to be designed in such a way, how can the method be validated? Similarly, what role would reliability play in an experiment set in a social setting? How can you recreate social settings for further empirical study?
I would love to read some examples of studies if you are familiar with any!
Many thanks.
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You asked about the validity and reliability. These are the critical points. If you chose your subject from very dissimilar groups and settings there is a greater chance to get reliable data as well as external validity which is important for researcher who want to repeat the experiment. Check this out if it gives you advice: https://pdfs.semanticscholar.org/6462/54a1698a66aacb6eceb66c126309c3311997.pdf
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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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What should be the minimum area for counducting a field trial?
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I think 25 square meter with 5 lines is good for /Control/Concentration. Its correct for chickpea, common bean......
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In a research having multiple treatments, say 4, there would be 4 treatment groups. In comparison, is one control group enough or should it have 4 control group.
Please note that all the groups here are randomised and homogenous.
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In the context of the main question of the discussion thread, control group may not be as large as treatment group.
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is it okay to get data from ansys or abaqus and build a model of neural network Or I must get the data from a realistic field experiment ?
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One can trust the results from simulation software as long as the inputs given to them were validated. The model setup in and results from ansys/abaqus needs to be validated against either experimental or analytical results. It could be an older study for which experimental results already exist or it could be one of the many cases you want to explore. Without proper validation, you MAY find a lot of numerical inaccuracies.
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For my master thesis i'm conducting a field experiment.
I have two dependent variables:
- The first is dichotomous (yes/no)
- The second is continious (number of seconds)
I only have 1 independent variable with three levels (categorical).
Now, i don't know which method is appropriate. Several topics have been opened about multiple DV's where i'm reading on MANOVA, multiple regression and canonical correlation.
Can anybody help which analysis is most appropriate? I will do my analysis in R.
Thank you in advance!
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Hello A. Kost,
The big question, for me, is whether it makes sense to look at the DVs simultaneously or separately. Unfortunately the description is too terse for me to offer a guess here.
If separately, logistic regression for the dichotomous DV (dummy coding the IV into two variates), and one-way anova for the continuous DV seem likely choices.
If simultaneously, canonical correlation is the most general, but a lot of folks seem to have difficulty interpreting results as easily as they do for tests of mean differences (whether for scalars or vectors, as in anova/manova). Here's a good guide for canonical presentations: Sherry & Hensen (2010), Conducting and interpreting canonical correlation analysis in personality research: A user-friendly primer. Journal of Personality Assessment, 84, 37-48. doi: 10.1207/s15327752jpa8401_09
Good luck with your work!
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My question is, How we can increase water hold capacity in open field experiment? What type of parameters we should concern more? Please refer some literature on it.
Highly appreciated in advance.
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Water holding capacity of the soil can be increased by adding organic matter and biochar to it. Please take a look at the following references.
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I analyze the microbial community composition from soil samples collected in an agricultural field experiment. Within this field, there are 3 plots (randomly distributed in the field) and for each plot, we collected soil samples at 2 depths and 3 different compartments (comp). We have a total of 18 samples. BUT I only have one observation per sample unit. So, basically the replicates are the plots. My first idea was to write the following model with adonis:
x ~ depth + comp
With plot factor considered as replicate. But I am not sure it's right because the plots are distant (several meters). They are not true replicate, right? Then, after I had discussed with some colleagues, it turned out that I should maybe use the following model:
x ~ plot/depth + plot/comp + plot
depth and comp being nested in plot. But in that case, I have no replicate at the lowest level, and it might be a problem for a permutation test... I don't know how to solve this problem. The experimental design is a bit tricky because we have no replicates at the lowest level. Therefore, I don't know how to deal with the statistic model.
If anyone can help me I would greatly appreciate :) Thanks for your time!
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Just to clarify on Igor's response; I think your advisors were probably trying to get at repeated measures rather than a nested design per se. Repeated measures is structurally the same as applying a blocking factor to the experimental unit (field). Nested design is similar to blocking, except that with a nested design your blocking factor is a treatment, and so you are interested in it's effect. Unless you are interested in the effect of your field choice, then I agree with Igor, it doesn't sound like your experiment is nested; it does however sound like it has repeated measures. Here's an example in R: https://www.r-bloggers.com/how-to-do-repeated-measures-anovas-in-r/
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I need to enter data in Hydrus 2D.
Solute was incorporated into the soil by field experiment applying urea and ammonium solution.
Results from laboratory are expressed in mg/kg and solution used in field in mg/L
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Are you asking about mg/L soil? If so you need the bulk density (specific gravity)
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I took light response curve data from my field experiment. QY and Amax
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Howdy Muhammad,
In principle a light response curve is a plot of net CO2 uptake rate (in µmol CO2 m-2 s-1) against different PAR or red actinic light PPFD's (in µmol photons.m-2.s-1 (at 625 nm for the red light of the Licor 6400).
You can then derive different parameters of photosynthesis from this type of plot. See for this - as an example -
Success,
Frank
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In what can best be described as a naturally occurring field experiment, it was noticed that soldiers who flew home directly from the 1982 Falklands war adapted less well than those who returned a week later by sea. Unfortunately, this is an anecdote. No actual measures of reverse-culture shock were taken. I'm wondering if the speed with which repatriates return affects their readjustment. Anyone know of any studies in which a period of decompression helps a major life-space transition of some type? Thanks in advance.
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Dear Uco,
after working as a surgeon in Tanzania for 2 years (accompanied by my wife and 3 children of 2,6 and ten years) we have all experienced a reverse-culture skock returning home to affluent Switzerland. For everyone in our family the problems were different: the youngest one missing his nanny and his first language Swahili, the older children missing their playmates and the freedom in nature. My wife had to adapt again to the circumstances of everything being available, of having no more support in childcare, kitchen and garden. For me as a surgeon the situation was more complicated: I was envied for my experience (2000 operations in 2 years), but at the same time suspicious ("you are no longer in bush!"). I am convinced we all would have profited from a decompressing period. Precondition would be to stay in a less affluent region (preferably with the same language as the definite destination). The question of financing this stay has to be solved, as no one earns enough money in a benevolent job. Maybe it should become part of the contract with the sending organisation.
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Is the artificial drought stress creating process same for pot experiment and field experiment?
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Dear Ms
you can find your answer on this article
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Any one have the method for calculate this parameter in field experiment. or this is just for wheat crop ?
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Convert the crop 1 yield to monetary terms (price x grain yield) and then adjust to crop 2 yield equivalent by dividing the crop 1 returns in monetary term by the price of crop 2 and then added to the crop 2 yield.
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Hello, Is it generally true that effect sizes are lower for ecologically valid field experiments compared with controlled lab experiments?
I'm not sure about this and I'm just trying to understand why the effect sizes in my field experiment was much smaller than my experimental self-report study, on the exact same IVs and DVs of course. Do you know anything about this? Some sources on this would help very much.
Many thanks for your help in advance!
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Hi, Pelin, usually the effect size or the minimal standardized difference we like to detect is given by the experimenter in adc´vance and from this the sample size is calculated to reach just this. When you do not design the experiment in advance then you have to live with the effect size coming out. In ecolocigal date the variance max be larger than in other fields so that the effec size for a fixed sample size is larger than in other fields. R-programs for all this you find in our book:
Rasch, D.; Pilz, J., Gebhardt, A. and Verdooren, R.L. (2011)
          Optimal Experimental Design with R,
          Boca Raton, Chapman and Hall,
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Suppose, I have 30 possible isolates of Azotobacter spp. I need to narrow it down to below five on the basis of their effect on rice vegetative growth ( maximum effect of an isolate in growth). If I want to go for traditional field experiment that will be a hell a large experiment and will take around five month to reach any conclusion (I want to do field experiment after narrowing it down to a feasible amount).
If anybody has any idea about in vitro experiment , less time consuming and effective, suggestions will be appreciated. Thanks.
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Eventual test of these micribial isolates is the field test where soil is inoculated with particular microbial isolate and the inoculated isolate if efficient , performs well while interacting with millions of other microbes which does not happen through in- vitro evaluation..
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I am interested in the building a rainout shelters to exclude precipitation for field experiments. 
Can anyone offer advice on materials, suppliers, and construction of shelters? I am interested in building about 48 units that look something like the shelters shown on this page:
As far as I can tell, these units are not commercially available as kits. If anyone has experience with building these, can you share your basic design and materials used? 
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We designed one:
250 sqM, 3 m heiıgh
weather move,
rain out,
Windows on 2 sides, 1 m above down and 1m above top,
two side doors, 2 door stye.
Use your local material
Good luck.
Nusret
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I have been carried out a field-experiment testing some criteria of vermicompost nutrient changes in vermicomposting. CV value was about 15-20%. Should i keep this data or test one more time? 
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Dear Vö,
CV% is dependent on some of those factors:
1. the experimental design,
2. the replication / plot size,
3. the experimental material you study.
The literature in your study area gives you a clue about the range of your acceptableCV%.
After checking the literatüre and you still think your CV % is higher than you wish to have, these are the things you can do:
1.Increase the number of replication and / or plot sizes,
2. Use a more effective expermental design, i.e. RCBD instead of RCB or triple lattice instead of RCBD, etc.
3. Use more unifom exprimental material, site etc.
Good luck.
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Hi all,
I started my PhD this year where I'm studying how hunting, parasites and vegetation influence the local distribution of ungulates. I'm using 2 species of antelopes that use similar niche (grazers) and live in an enclosure of 4600 ha with no natural predators. I would like to collar 10 females per species with gps devices, but I know these devices are expensive and I don't have the budget for buying 20 devices.
I want to get information from 4 locations (early morning, noon, afternoon and evening) per day during a period of 6 months. I also want to take the data from the devices with out capturing the animal (transmission of data to cell phone or other gps or drop-off system?), because  I will not be able to catch the animals again after collaring them.
 Do you know anyone who can give me advice on how to build or where to buy these devices with a limited budget?
I checked Mataki project (http://www.mataki.org) and I contacted people from Blake Allan paper (https://www.researchgate.net/publication/277385807_A_cost-effective_and_informative_method_of_GPS_tracking_wildlife ) but I would like to know if you have more ideas.
thank you
regards
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Thank you very much Jacob
Have a nice day,
Chloé
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Dear colleges,
I was conducted two field experiments in two consecutive years in order to estimate genetic diversity analysis among forty fourth genotypes of lentil (lens culinaris M.). Morphological characterization was done based on different parameters like plant height, pod per plant, seed per plant, 100 seeds weight… The experiments were carried out following an Alpha Lattice design with fourth replicates. Data were recorded from both years. I would like to know if generalized linear mixed model (GLMM) is applicable in this case, considering a genotype as a fixed effect and a bloc as random effect.
Thank you for your response.
Best Regards,
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Dear Djouher,
I agree with Salvador. If using GLMM, that would be more robust since the data may not be normally distributed. Actually, I would not suggest using ANOVA in any situation, since ANOVA requires too many strong assumptions, such as homoskedesticity, which is very hard to verify. In my opinion, ANOVA's only advantage is that it is easier to be computed by hands, but in this era, who needs to do empirical studies by hands?
Hope it would be any help to you.
Sincerely,
Kuan-wei
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How to analyze the data collected?
Can active search of Naja kaouthia in nests or rat hole be used to estimate population size? 
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I'm shopping for field photosynthesis measuring systems.  The LiCOR 6400 and PP Systems CIRAS-3 both have the option of estimating apparent electron transport rate (ETR) and PSII yield based on fluorometry: basically looking at Chlorophyll fluorescence. The WALZ PAM 2500 is exclusively a fluorometer.  In contrast, the LiCOR 6400 and PP SYSTEMS CIRAS-3 are mainly estimating photosynthetic rate based on differences in CO2 and H2O vapor.  
Have people tried both fluorometry and gas exchange approaches to estimating photosynthesis? What are the relative advantages of both approaches?  Do you really need both systems to get an estimate of photosynthetic rates?  What are the relative errors?
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Marcin is mostly right, but you should note that fluorometry can treat more leaves per time. So it might be the method of choice for screening of larger number of plants. 
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We had several samples losses of Barber pitfall traps in our Alpine sites due to marmots that were attracted by the conserving solutions or trampling cattle. We used ethylene glycol, but we switched to the non-toxic propylene glycol, unfortunately both have a sweet taste, the latter with reported with less attractiveness.
I wanted to ask the community, if there are tips for metal cages that cover the pitfall traps.
I already designed some prototypes (I love to tinker :-) ) and ideally the cages should:
- not influence the catch pattern and rate
- be robust but also of light weight
- not be too expensive
Thank you,
Best, Michael
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Make any pitfall solution (antifreeze, saltwater) distasteful to mammals by adding a little formalin (not enough to harm insect integument for short periods).
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I want to create range of light intensities in the 2×5 m plots under field condition. Which materials can be suitable for this purpose?.
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Dr. Ali Reza Yousefi,
I consider that there is no best way to change light intensity under field conditions. There are various materials to do this like shading nets and screens as already pointed out by Dr. Bongi and Dr. Graham, respectively. I agree strongly with Dr. Graham if the point is the modification of light intensity. The suitability of each available material in field conditions depends primarily on its resistance to extreme weather conditions e.g. hail. In an open field, another problem is the greater variability of a number of variables, in general, in relation to a glasshouse and the effective control of this variability.
The attached publication, with regard to shading by shading nets, may be useful for your research.
Best Regards
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Dear all,
             Im working in a field experiment with 20 traps (pitfall traps to collect ground atrhopods) in a treatment field and 20 traps in a reference field (so potentially spatially autocorrelated).
I performed a nMDS (non param multidimensional scaling) plot to assess multivariate ordination of those samples and I plotted also 95% confidence ellipses to visualize effective discrimination between the treatment and the reference field. Then I would like to have a statistical measure of this discrimination so my idea was to perform a perMANOVA (adonis function in R software) to test dissimilarity between fields. So my question is: 
-Can I use perMANOVA with such experimental design? If not, is there a way to deal with such autocorrelation? Suggestion on alternatives?
Thanks a lot
Alessandro
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Thanks Remi, I will try to make a Mantel test comparing animal abundance distance matrix with the location (of the traps in the field) matrix. I should solve the problem in this way. Thanks a lot
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For purposes of scientific reviews and to obtain a better sense of the state of biochar implementation outside of broad-field agriculture, I am interested in learning about any prior or ongoing field experiments on biochar effects on plant productivity, soil processes, and biogeochemistry.  A brief "meta-data" description would be valuable (location; year implemented; size of treated areas; biochar dosage(s); plot replication; type of biochar used including feedstock, pyrolysis method, and peak temperature; soil type, including texture and pH; ecosystem or target plant species examined).
Depending on responses, I would consider collaboration on a review with contributors.
I have attached a file with my lab's ongoing field trials as a template.
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I am looking for a small mobile unit that could be used to perform DNA extraction, PCR and gel or capillary electrophoresis in the field, i.e. Not in laboratory conditions. The purpose of the kit is to perform species or individual ID. Samples would range from muscle to hair and PCR would use mtDNA or STR primers. 
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Hi Olutolani Smith
I know but not use about Palm PCR gadget
All the best,
Vladimir
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I am basically an Agribusiness professional, interested in Random Field Experiment projects. Kindly help me to start with this
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Dear Anderson,
I am ready with my draft proposal. Kindly, provide your email id so that I can send the same for your review.
Dr. Rams
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We would like to observe the development of single insects on plants in the field. The insects usually settle on the bottom side of the leaf. Leaves are quite large (about palm-size) and more or less waved with some strong veins. The experiment will run about one month during summer, and therefore possible physical damage by the cages, rains and wind are additional challenges (the standard setup with metal clips and sticks can be problematic, since it is unstable and susceptible to wind especially with upper leaves which are about 30-50 cm above ground).
Now we are unsure what type of caging to use and a few ideas we had so far are:
  • Light-weight clip-cages without sticks (still fragile setup, mounting only possible at leaf edges in larger leaves bec of clips)
  • Gauze nets (covers whole leaf; prone to getting wet during rain, to heavy then or insects might drown)
  • Perforated plastic bags (too high humidity and temperatures within bag)
Do you have any better setup for such caging or another cage type or an idea how to mount a clip cage safely on such leaves in the field?
Thanks for any input.
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Any cage will modify the microenvironment about the insect. It will also influence the physiology of the leaf on a spatial scale relevant to the insect. The problems you mention for gauze nets and perforated plastic bags apply to all cages, are unavoidable, but may be worse for some strategies.
How about using a flat magnet to attach the cage to the leaf. I would make something like a clip cage, but glue the magnet to the base and use a magnet of opposite polarity on the other side (or maybe something like iron foil). It will be fairly heavy, so attach a bamboo skewer to the branch in such a way as to provide support. Might use thick wire to do the same thing.
How about using the clip cages and "gluing" them on with something like tanglefoot. I would use the bamboo skewer idea to add support. I tried to think of a non-toxic glue that would work and didn't have any ideas. Bioquip sells some nice small clip cages that may work.
In part, how well this works depends on how strong the wave is in the leaf. You could buy foam pipe, and cut it to match the wave in the leaf. Then use tanglefoot.
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We have conducted an experiment to check the reliability of postion, velocity, and altitude as obtained by GPS tracking data (see attachment). We did an extensive search of the literature without finding much. Anybody aware of papers concerning extensive, experimental testing of the reliability?
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 @Graham: Thanks, I have been away from work for sometime and just saw your posted response. I will take a look at the papers you suggest.
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We are trying to learn more about any kind of species interactions (competition, predation, disease transmission etc.) between native and non-native (invasive) crayfish in the eastern United States. We are designing some laboratory and field experiments to assess the potential impacts of invasive on natives (focusing on Virginia), so any additional information would help us do a better job with new research program in my lab. Thanks.
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I would contact your local Army Corps of Engineers Biologist. The Corps do regular biodiversity assessments of the properties they manage, which includes a lot of streams and marshlands. They will sometimes even do complete watershed assessments. They would likely be able to tell you if they have seen any invasions and resultant changes in species profiles. That could give you an idea of relative competitive fitness. It wouldn't be a published study or experimental results, but could lend itself to some important observations.
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I want to use a mobile eye tracker for some real world experiments and still trying to search for the best option for it. Any opinions or experiences about this device?
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Hi, I used Positive Science eyetrackers when I was a Products Researcher at Procter & Gamble. It's the best mobile eye tracker on the market if keeping every eye tracking movie as a usable data source is important to you. It's the most flexible & has the best tools for getting an eye tracker to work, once you get the hang of the software. In that arena, it easily outperforms systems that are 3-4 times more expensive. I've used Tobii, ASL, SMI, and Eyelink, and the Positive Science Ultraflex is the absolute best at delivering accurate tracking in a wide variety of conditions. I've used Positive Science eyetrackers in grocery stores, in consumers' homes, even in rural india - it works everywhere.
Coding data is really the big problem for all head mounted eye trackers. There are some eye trackers that have provided coding solutions, (dikablis, Tobii, ASL) but those have largely come at an insane cost, and they also deliver a terrible track. The Dikablis,. for example, costs $60,000 and puts its scene camera in the middle of the forehead, unnecessarily increasing parallax error. The Tobii glasses just plain don't work about 70% of the time. When they do actually work, and when you have put their markers on everything, it's OK, i guess, but you never really know when they're working until after you're done, and they also cost $45k. (last i checked, which was a few years ago). Also, if someone isn't tracking in a Tobii, it's a complete black box, so you just have to give up & try a new person. ASL has a reasonably good eye tracker, and they are doing a lot to improve their motion matching algorithms, but there are still risks where you can lose the entire video if someone moves their head too fast & smears the image (from what I can recall, I last tested their technology in 2010)
For full disclosure, I worked with Positive Science and with the Rochester Institute of Technology to develop their coding solution, Semanticode. This is mostly because the other solutions on the market were so bad. I've divested all financial interests and receive no money from the licensing of the software, however. But I'll freely admit bias when it comes to the hand-coding solution - I think it is the best way to code fixations for real-world eye tracking experiments, mostly because you still need humans to categorize fixations, unless you're running Tannenhaus-esque visual world experiments, or something like that.
So, just one opinion, but I think the Positive Science is best in class at delivering an accurate track (if you are willing to learn the extensive tools that they have for tracking gaze), and that it has the most accurate coding solution, given that the other trackers (like Tobii and Dikablis) are so ungodly terrible at, well, eye tracking. SMI had an interesting moveable ROI tool in their software, but their hardware are also high in cost, and you probably aren't saving yourself a lot of time by coding videos that way vs. coding them in SemantiCode. Positive Science way outperforms SMI on cost.
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When we use the dirac delta function to solve the paradox of divergence of a 1/r^2 field, we say that divergence of the field is infinity at r=0 as delta function reaches infinity at r=0 which means that there is infinite flux passing through a minute volume. But since it can be mathematically proven that a point charge is nothing but a uniformly charged solid sphere with a very small radius, the electrostatic field at r=0 is 0, which contradicts the result obtained using dirac delta function. Why?
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First, the field going as 1/r^2 (and naturally diverging at r=0), corresponds to
a classical model and describes the field in the vacuum, that is, outside the point charge.
Second, the Dirac's delta is not a function but a distribution. Distributions have such a 'bizarre' behavior because they are defined to be finite under integration.
Finally, that is not true that a point charge is exactly an sphere with small radius. The issue here is precisely, that a point charge is a mathematical point, that is, has no measure. That is precisely why we describe point charge densities by using Dirac's deltas: to have finite fluxes! So there are no paradoxes. It is just a convenient description to treat classically objects (particles and fields) that are quantum in their very essence.
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Looking to freeze down biopsies in the field, collected from rare wildlife species. A Mr Frosty works well in a lab set-up, using a -80C freezer, but not suitable for field work. A Liquid Nitrogen flask can be taken into the field. Any ideas as to which company or supplier may have such an apparatus available?
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Wow.....
Much appreciated...
Dr Paul Bartels
NZG Biobank
National Zoological Gardens of SA
Boom street, Pretoria. 0001. South Africa.
Tel: 082 990-3533 (..27 82 990-3533)
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I am looking for a possibility to survey insects in the field using video cameras, possibly also at night. Does anybody have some good or bad experience with this kind of survey and some indications about possible material to use?
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Before we can make any recommendation about UV or other filters, it would be helpful to know what you mean by "survey." Do you mean that you want to non-destructively "census" insects at a feeding stations or a light source (or a white sheet with a light behind it)? That is, are you wanting an automatic camera taking shots at regular time intervals? Or do you mean that you are walking transects, using spotlights, and then taking photographs?
Some thoughts:
- If using simple spotlighting to census insects (at night) and then repeating the transects during the day, a good wide-angle lens will no doubt be necessary. For small insects, you may want to build a rig with a reverse-mounted lens with a flash diffuser (on a bracket). (To save money, don't buy a macro lens. Just take your regular 18-55mm--or shorter--lens and turn it around. You need a cheap mount adapter to do this.) You can add to this setup using extension tubes.
- For daytime shots, you may avoid getting close to your subjects altogether. I suggest trying a larger lens (say, a 70-300mm) with a set of extension tubes. You'll be able to stay far enough away to prevent insects from flying but get high enough magnification to capture the insect well.
- Concerning UV lens (@Madan Gautam) : You probably don't need this if you just want to identify the insects. If you do want to get into animal visual modeling (there's a great new literature out there for using standard DSLR cameras for detecting UV), be warned that this is an expensive prospect. The associated equipment can be very expensive. One of the standard ones (Nikon Nikkor UV lens) runs about US$8000. Of course, the lens is only the beginning. Because the camera sensor elements sensitive to UV are also the ones sensitive to other dominant wavelengths (in the human visual range), it is necessary to block out the visual-range wavelengths. So, you need a UV-pass filter. Of course, because you want to discriminate between UV and visual, you also need a filter that cuts out UV and isolates the visual range. These filters are expensive. So, let's hope you're only doing this for survey purposes.
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I measure Net Ecosystem Exchange in peatlands and I'm having problems with the temperature inside the chamber. Right from the start, it increases dramatically. Such an artefact affects photosynthesis, which makes the measurement unrealistic and, thus, unexploitable. Has anyone been through such a problem already? Any idea how to resolve it? Any references about this problem?
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Here you find two papers addressing the issue:
Laine et al., 2006. Estimating net ecosystem exchange in a patterned ecosystem: example from blanket bog Agricultural and Forest Meteorology, 138 (2006), pp. 231–243.
Anna Laine, Terhi Riutta, Sari Juutinen, Minna Väliranta, Eeva-Stiina Tuittila. 2006. Acknowledging the spatial heterogeneity in modelling/reconstructing carbon dioxide exchange in a northern aapa mire. Ecological Modelling 220 (20): 2646–2655.
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Experiments could be laboratory based and theoretical in nature, or applied field experiments that provide valuable cross-site comparisons, could be economic, ecological, psychological, or all of the above. Experiments could be large scale, centralized projects (such as a "Biosphere II" approach) or smaller precision research tools.
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Timothy, ERoEI as a key to the determination of whether or not there is a sustainable energy source to replace fossil fuel technology , which is not sustainable for other reasons, is discussed in the paper http://dematerialism.net/eroeistar.htm and my new/old blog at http://eroei.blogspot.com/ .