tropical ecology, assessment and monitoring (team) …

TROPICAL ECOLOGY, ASSESSMENT, AND MONITORING

(TEAM) INITIATIVE

ANT MONITORING PROTOCOL

DRAFT PLEASE DO NOT DISTRIBUTE!

TEAM Initiative Members Gustavo Fonseca, Ph.D. Senior VP for Science and Executive Director, CABS Thomas E. Lacher, Jr., Ph.D., Senior Director Puja Batra, Ph.D., Project Director James Sanderson, Ph.D., Research Scientist Scott Brandes, Ph.D., Post-doctoral Researcher Alvaro Espinel, Manager, Database Programs Caroline Kuebler, Project Coordinator Ariel Bailey, Administrative Assistant James Heath, Program Associate

TEAM Ant Protocol Draft do not distribute

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Introduction

Ants constitute a staggeringly high proportion of the earth’s biomass (Fittkau and Klinge 1973) and species diversity (Erwin 1989; Holldobler and Wilson 1990). They provide a number of crucial services important to overall functioning of ecosystems, such as soil enrichment and turning, seed dispersal, control of herbivorous insects, symbiotic interactions (Holldobler and Wilson 1990 and references therein), and have been the subject of a vast amount of research over several decades which has increased our understanding of ecology, evolutionary biology, animal behavior, and the list goes on. Thus, a multi-taxon monitoring program that aims to capture change in elements known to be critical to overall biodiversity and ecosystem health must necessarily include ants. In addition to the potential value of ants as a surrogate group for overall diversity (see (Alonso 2000) for a detailed discussion), they have great promise as indicators of ecological change (Nepstad 1995). Ant community structure has been shown to be sensitive to heterogeneity in a number of environmental factors. For example, data from both within and across site studies have shown relationships between ground dwelling ant communities and soil properties (Catangui et al. 1996; Bandeira and Harada 1998; Bestelmeyer and Wiens 2001), seasonality and climate factors (Bestelmeyer and Wiens 1996; Feener and Schupp 1998; Kaspari and Weiser 2000), vegetation density (Bestelmeyer and Wiens 2001), vegetation communities (Majer and Delabie 1994; Majer et al. 1997; Morrison 1998; Bestelmeyer and Wiens 2001), and plant productivity (Kaspari et al. 2000; Kaspari et al. 2000). Many of these factors have been shown to be currently undergoing measurable changes due to climate and land use change (Root and Schneider 2002; Walther et al. 2002; Parmesan and Yohe 2003), and these effects are predicted to continue into the foreseeable future (Bazzaz 1998; Enquist 2002). Many aspects of these factors will be monitored by the TEAM Initiative. Thus, the correlation of multiple data sets taken in the same areas over the same time span will allow not only intrasite correlations between ant communities and various biological, climatic, and physio-chemicalfactors, but also will allow for detection of intersite trends that may be occuring regionally. Furthermore, by monitoring communities of ants over time, we will add enormously to our understanding of the baseline levels of fluctuations in ant diversity and turnover at different spatial scales. Specific questions By monitoring leaf litter ants several times per year, we will address the following questions:

1) What is the baseline level of fluctuation in leaf litter ant communities in tropical forests? How does that baseline differ across regions?

2) Are there trends in indices of species richness or relative abundance over time? If so, do those trends correlate with trends in climate measures, soil properties, tree growth rates, phenology/litterfall, or measures of land-use change? At what spatial scale do trends occur over? Are similar trends occurring across regions?

3) Is community composition changing over time? If so, in what way? (e.g., Are species being lost or gained? Are there invasions of exotics occurring? Is community structure, i.e., dominance and sub-dominance shifting? Are species which are relatively restricted in geographic range more vulnerable to population changes?)

4) At what spatial scale does turnover in litter ant community composition occur? Methods for monitoring ants Because of their ubiquity and high species richness, there are many techniques and microhabitats useful for ant collection. These include sampling from the leaf litter and/or soil using a variety of extractor types, using pitfall traps to collect ants from the soil surface, setting baits to attract various foraging guilds, canopy fogging, beating vegetation to collect ants in the shrub layer, hand searching, and still others. The utility of these various techniques depends entirely on the questions being addressed, and results


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quantifying aspects of diversity may or may not be comparable across sites due to differences in collector experience, uneven presence of certain guilds, etc. For the purposes of TEAM monitoring, the technique chosen must provide a method which minimizes the effects of varying collector experience, maximizes efficiency in species: effort ratio while also reliably capturing ants within the same species guild. In a massive study conducted in the Atlantic forest region of Brazil which compared the efficiency in capturing species richness of 17 different ant collection methods, Delabie, et al ( 2000) concluded that the combination of pitfall traps and leaf litter sampling using Winkler extractors captured 50% of the total species richness found. A standard protocol was agreed upon by many members of the myrmecologist community (Agosti et al. 2000), and this protocol, known as the Ants of the Leaf Litter (ALL) method is rapidly becoming the preferred method used around the world for inventory and monitoring of ants. TEAM monitoring will utilize the ALL method, with some minor modifications in transect length and spatial design. The modifications will not alter the comparability of TEAM data to other studies examining species richness and abundance, since such estimates are typically made on a per sample basis, and not a per transect basis. TEAM monitoring using ALL will be conducted at a massive scale, sampling four transects of 100m length in each Integrated Monitoring Array (IMA) four times per year.

Spatial layout The ant transects will run along side soil termite transects, thus this layout was designed to accommodate both types. The sampling design for leaf litter ants consists of replicated 100 meter transects, with sampling stations spaced at ten meter intervals along the transect. Each sampling season, four 100 m transects will be placed inside the 1km2 Integrated Monitoring Array (IMA) (figure 1). For the purposes of placement of ant (and termite) transects, the IMA can be considered to be divided into four quadrants of equal area, with invisible lines running down the central horizontal and vertical axes. One transect will be randomly placed within each of these quadrants. Constraining the placement of each transect’s potential location to only one fourth of the IMA will allow for some degree of randomization while also achieving even coverage of the array over time, thus maximizing the chance of capturing some of the heterogeneity of microenvironments that the large IMA contains. Transects will be placed in the grid square between the pair of (x,y) coordinates chosen and the next higher number of each. For example, if the (x,y) coordinates chosen randomly are (04, 600), the transect will be placed in the grid square that falls between points (04, 600) and (05, 700). The divisions between quadrants run along line 06 and line 500 (figure 1.) Using a random number table, four sets of randomly chosen (x,y) coordinates must be selected, with the coordinates falling between the following numbers:

01-05, 000-400 01-05, 500-900 06-10, 000-400 06-10, 500-900

Once a grid square is used, it will never be re-used until all possible locations have been used. Thus, after the first season of sampling, it will be necessary to carry a list in the field of all the locations which have been previously used and are therefore, not repeatable. If the grid square selected has previously been sampled, random number selection must be repeated until an appropriate square is found. If there is standing water in the middle of the grid square, shift anywhere within the same square to a dry area so that the 100 meter transect can be placed perpendicular to the trail. If the amount or distribution of standing water excludes the possibility of placingthe transect anywhere in the grid square, repeat the random number selection process until a dry grid square is found. If all available choices (i.e., all non-repeatable locations) have standing water in them, make a note of this in your notebook, and select a dry repeat location. Similarly, if you reach the grid square and find that it is being swarmed by army ants, make a note of it and repeat the selection process. The reason for this is not simply to avoid sampling an overabundance of army ants, but more to facilitate the field work in the leaf litter without the obvious problems to collectors that army ants might cause.


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The transect should run such that it starts along a major trail, i.e., perpendicular to the trail, halfway (50 m) between the two corners. The first sampling station should be inside the grid square, 10 meters from the trail, so that the last sampling station falls at the edge of the grid square where there may or may not be a temporary maintenance trail running between IMA numbering stakes. To avoid confusion, flagging for different groups of TEAM variables is color coded. All insect sampling locations and/or entrance points from the trail should be marked with blue flagging only.

Equipment

(see Appendix 1 for a list of equipment suppliers) For litter sampling (per transect): random number table sighting compass 100 m nylon rope knotted at 10 m intervals blue flagging 1 1m2 quadrat made of wood or PVC (movable joints; one corner should be openable) 1 machete 2 pair gloves 1 small hand rake 1 small hand trowel 1 watch with seconds or stopwatch 1 litter sifter 10 litter stuff sacks (included with the mini-Winkler apparatus) and some extras 2 waterproof notebooks pencils large plastic bag, large enough to carry 10 mini-Winkler samples without compressing them For pitfall traps (per transect): soil auger (or sharp edged pipe with diameter of the pitfall cups) to dig pitfall trap holes; must be strong enough to cut through roots. 10 durable plastic cups, 7-8 cm diameter, 8-10 cm depth, with no ridges inside or at mouth 1 liter 75% Ethanol 5 ml liquid detergent 10 small plastic plates, few cm larger than diameter of cups; used as a roof for the pitfall trap thin wooden skewers labels (card stock) squeeze bottle forceps waterproof and Ethanol proof ink pen For measuring ecological conditions Kestrel 3000 Pocket Weather Station Electronic soil probes for pH, temperature, humidity Small ruler (15 cm) For processing mini-Winkler litter samples and pitfall trap contents (per transect): 2 pieces nylon rope (5 m) 1 2x3 m plastic sheet (only 2 are needed in total) 10 mini-Winkler apparatus (each mini-Winkler contains a mesh bag, and the external cloth portion) and a few extra 2 light colored plastic trays medium width (5 cm) paintbrush


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fine (2-3 mm) paintbrush forceps 10 plastic cups with no ridges inside or at mouth 2 liters 95% Ethanol (source alcohol, not denatured EtOH) squirt bottle 10 vials (25-50 ml) with tight caps 1 large ziploc bag labels (card stock) waterproof and EtOH proof ink pen scissors

Methods Field sampling Running the transect Use a compass and a 100m nylon rope that has already been measured and marked with knots at 10 m intervals. The first sampling station will be 10 m from the trail so tie the rope to a tree just inside the grid off the trail. With the rope to help you run a straight line, sight intermediate landmarks through the sighting compass that fall in the desired direction. The heterogeneity of microhabitats is important in maintaining species diversity, so do not avoid any types of microhabitats that fall in the straight line, even though some may be difficult to sample in. Place a piece of blue flagging at every 10 m increment until you have placed 10 such flags, all in a straight line. It is best to leave the rope in place, tied to trees at either end so that you can easily find the next point without the need to search for the flagging. If there are trees, fallen logs, etc in the line, the transect must include them. The knots (and flagging) will be the points at which you will sample leaf litter and place pitfall traps. Leaf litter samples (modified, but largely after Bestelmeyer et al. (2000)) The leaf litter sampling is to be done when the litter is somewhat damp, but not saturated. Wait a few hours after a heavy rain to collect litter. Transects are best done with two people working on the litter samples and one person simultaneously placing the pitfall traps. Hereafter, the terms “Winkler” and “mini-Winkler” are interchangeable. Try as much as possible to walk on the side of the rope that will not be sampled, to avoid trampling or disturbing the litter. At each flagged location on the transect, place the 1 m2 quadrat on the ground. To ensure that your bias of having seen the entire transect while running the line does not influence where you place the quadrat, decide in advance of running the transect line where relative to the 10m marks you will place the 1m2 litter sampling quadrat. For example, decide on the rule that you will always place the quadrat on the right side of the line, with the knot on the rope falling in the bottom corner of the quadrat. A rule against this type of bias will result in being able to sample many of the microhabitats that you may encounter on the transect such as treefall gaps, fallen logs, etc. The quadrat has one corner left open so that it can be put around trees and shrubs that will fall in the sampling spot. If there are trees too large to put the quadrat around, shift the quadrat so that one edge is touching the tree trunk. If there is a fallen tree on the ground, include it if possible since it will probably have an accumulation of litter on it. If it is too large, place the quadrat as close to it as possible so that one edge is touching the fallen log.

It is highly advisable to wear gloves while handling the leaf litter. Remove large sticks in the quadrat. Use a machete to chop open decaying logs inside the quadrat. Scoop litter inside the quadrat toward the center, using the hand trowel and hand rake (figure 2). Break open twigs and clods of soil. Tie off the bottom of sifter, and place litter in top of sifter up to ¾ full. Break up any more twigs or pieces of rotting wood that are in the sifter. Hold both handles and shake the sifter in all directions continuously for 30 seconds (figure


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3). This shakes the smaller litter and invertebrates in it to the bottom part. Thoroughly turn the litter over by hand and shake for another 30 seconds. Mark the cloth stuff sack with a label with the following information, written in waterproof and alcohol proof ink:

Date: mm/dd/yr Array: 1-6 Transect grid square coordinates: (x,y) Sample number (1-10) Winkler

Place the litter stuff sack below the sifter sack, and untie the sifter sack. Shake all contents of sifter into the litter stuff sack (figure 3), shaking the cloth to dislodge any insects crawling up the sides of the sifter. Close the sample bag tightly. When carrying the litter samples back to the field station in a large plastic bag, take care not to compress them or place anything on top of them. After returning to the field station, the samples will be placed into mini-Winkler sacks and left for 48 hours during which time the ants will migrate out of the leaf litter into a collection cup or bag. This process is described in a later section. Pitfall traps Pitfall traps will be placed along the same transect and at the same time as the litter sample collection, and left in place for 48 hours. Place pitfall traps in a consistent place relative to the quadrat, about 10-15 cm in front of the litter quadrat, in approximately the midpoint of the front edge of the quadrat. Using a soil auger or metal pipe whose diameter is roughly the same as the plastic cups, dig a hole that is the same depth as the cup. You will have to cut through roots in many instances. If the roots are impossible to cut through, shift the location of the hole by a few centimeters. If possible, cut the hole without removing the litter. This will make it easy to use the edge of the hole to measure litter depth (described below). Place a plastic cup in the hole, flush with the ground (figure 4). If the hole is too wide, you will need to pack in soil around the sides to keep it in place. In order to keep soil out of the cup (which gets into the sample and is difficult to remove during sample processing), stack a second cup inside the one that will be the trap while you arrange the soil around it. Also rearrange the leaf litter so that it comes up to the edge of the pitfall trap. On a label, using waterproof and alcohol proof ink, record the following information:

Date: dd/mm/yr Array number: 1-6 Transect grid square coordinates: (x,y) Sample number: 1-10 Pitfall

Place the label in the cup and pour in about 2-3 cm depth of 75% ethanol that has been mixed with a few drops of liquid dish detergent (about 5 ml detergent per liter). The detergent breaks the surface tension so that insects which fall into the trap actually sink into the liquid, and also prevents rapid evaporation of the ethanol during the 48 hours that the trap is left in the field. Cover the cup with a “roof” that is held a few centimeters above the cup. This can be made of easily available materials, such as a small plastic plate held up with thin wooden skewers (figure 4). The roof should be only slightly larger in diameter than the cup. Leave traps in place for 48 hours. Collect them on day 3 by transferring the contents of the trap to a 50 ml tube with the aid of a squeeze bottle of 75% ethanol and forceps, putting a label inside the tube, and closing the tube tightly. Any labeling information that you write on the outside of the tube is likely to be erased if the alcohol drips or spills on it, so be sure there is a clearly written label inside the tube, preferably positioned so that it can be read from the outside of the tube.


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Ecological data While collecting litter samples and placing pitfall traps, it is crucial to collect some basic data on ecological conditions on the transect. Appendix 2 contains a copy of the data sheet that must be filled in completely when a new transect is sampled. When starting the sampling, using the Kestrel 3000 Pocket Weather Station, record the air temperature and relative humidity. In three 10 m x 1m sections along the transect, collect data on the vegetation, canopy, ground habitat, and leaf litter. The ten-meter sections where you will collect these data are in from meters 1-10, meters 40-50, and meters 90-100 along the transect line in a 1m wide band. The vegetation description is a quantification of the number of live stems in each of the 10m2 area sections. Count and record the number of live stems in three different height categories and one girth category:

< 1m height < 2 m height >2 m height > 10cm DBH

Within any given section, if trees land in partially inside and partially outside the 10m2 area, alternate whether you count it or not, i.e., count the first such tree in your description, exclude the second such tree, and do on. For canopy description, estimate the height in meters of the main canopy layer and the emergent layer. Qualitatively describe the canopy cover as one of the following categories: Closed

Partially open Open with sub-canopy secondary growth Open with very little or no sub-canopy growth (new gap) For ground habitat description, estimate in increments of 10% the amount of ground cover of each of the following categories: Bare soil Stone Live vegetation, including roots Decomposing wood Leaf litter In this case, ground cover refers literally to what is touching the ground. For example, if there is a treefall whose trunk is not touching the ground, and under the trunk there is bare soil, the relevant category would be bare soil. In cases where a category is represented by less than 10% but more than zero, record 1% as a way to indicate its presence in very low relative amounts. Leaf litter conditions also need to be recorded, as they are important in determining the immediate environment that the ants experience. Using electronic soil probes, record the following: Soil pH Soil temperature Soil humidity Also measure the litter depth in cm with a small ruler. The easiest way to do this is from the hole that you have dug for the pitfall trap, if you are able to cut the hole without disturbing the litter around it. Extraction of ants from litter samples using Mini-Winkler apparatus The following can be done in the field under a large tarp in a place that is protected from the wind and rain. However, it is much easier if you bring the samples back to the field station. This process should be done on the same day as the samples were collected in the field, and will be left in place for 48 hours. It is very useful to make a note of which species of ants may be foraging in the room where you are working. They are often invasives or extremely common ants that can be easily identified (W. Overal, pers.comm.) At some later date, this information may be useful in discerning a contamination problem. If you are working with the samples in a field shelter and not an enclosed building, you will need to be extremely vigilant about making sure that ants do not crawl on the ground sheet on which you are working. They will be


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easily confused with escapees from your sample otherwise, and you will contaminate the samples with ants that did not come from your transect. Hang ten mini-Winklers from a rope strung horizontally in an area of a room where they will not be moved, blown by the wind, or disturbed (figure 6). Tie an empty cup in the bottom of each. It will later be filled with ethanol. In most instances, one mesh sack will be sufficient to hold a single litter sample. However, in the event that the litter from one sample does not fit entirely in one mesh sack, it will be useful to have a few extra mini-Winklers on hand. Place a large plastic sheet on the ground. From now on, do all work on it so that escaping ants can be seen and captured. Working above a plastic tray to catch litter that falls through the mesh, transfer the litter sample from a single litter stuff sack to a mesh inlet sack. Use the 5 cm width paintbrush to collect the litter that has fallen in the tray and transfer it back into the mesh sack (figure 5). You will have to repeat this process several times to get all of the fine litter particles in. If ants escape during this process, pick them up with either forceps or a fine paintbrush moistened with alcohol or water and put them into a vial or back into the litter. Periodically shake the mesh sack to let the contents settle and to remove air pockets. Do not leave any material in the tray or on the ground sheet. The mesh sack should remain flat even with the litter in it in order for it to be effective, so do not stuff it so full that it bulges. Between samples, be sure to clean the brushes and trays of any remaining debris particles, so that tiny ants which may be lodged in them do not end up in the wrong sample. Carefully hang the mesh sack inside the external mini-Winkler sack (figure 6). The mesh sack should not touch the walls of the mini-Winkler sack. If any debris falls into the cup, empty the cup back into the mesh sack. Partially fill the cup with 95% ethanol and tie this to the bottom of the mini-Winkler. Put the sample label in the cup, and also transfer into the cup any ants that you caught while transferring the litter. The ants will fall out of the mesh sack into the alcohol filled cup. After 24 hours, each mesh sack must be carefully taken out, emptied into a plastic tray and put back into the mesh sack again, following the same procedure outlined above. This process stirs up the litter and allows more ants to be collected. Leave the mini-Winkler apparatus in place for another 24 hours. At the end of the period, remove the cup, pour the contents of it into a 50 ml plastic tube, rinsing cup with EtOH. Be sure to include label with the sample. Close the tube tightly. Keep all tubes from a transect together in a large ziploc bag, also labeled. Personnel needed for field work and sampling rotation The entire process of running the transect line collecting and sifting 10 litter samples, placing 10 pitfall traps, and taking ecological data takes a team of four people about two hours, after the initial process has been learned. Two transects can be done in a single day. Transferring 10 litter samples to mini-Winkler extractors takes two people about two hours also. Thus, a typical day in the field and lab will take a team of four people 8-10 hours, including travel time to the arrays, grid coordinate selection, and walking to the grid locations. Since transects need to be revisited to pick up the pitfall traps 48 hours after the transect is run, the most efficient way to arrange sampling of different arrays is to do it in such a way as to minimize the number of trips to a single array. The following chart illustrates an efficient sampling rotation.


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IMA # transect # Day 1 Day 2 Day 3 Day 4 Day 5 (etc) 1 1 Transect Pick up pitfalls 1 2 Transect Pick up pitfalls 2 1 Transect Pick up pitfalls 2 2 Transect Pick up pitfalls 1 3 Transect Pick up pitfalls 1 4 Transect Pick up pitfalls 2 3 Transect 2 4 Transect 3 1 Transect 3 (etc) 2 Transect Here, on days 1, 2, 3, and 4 only one array is visited. On day 5, two arrays are visited, and so on. In this rotation, starting to sample transects in IMA 2 on day 2, before all transects on IMA 1 have been done prevents an extra visit to IMA 1 since you can pick up pitfalls from transects 1 and 2 on IMA 1 at the same time that you sample transects 3 and 4. Sampling will be done four times a year: early rainy season, peak rainy season, early dry season, and peak dry season. Specimen processing and identification Samples should be identified by an expert taxonomist, or under the supervision of one. Details regarding this process will be specified in later versions of this document.

Field data forms, database forms, and data entry A sample data sheet for collection of ecological data on the transects is included in Appendix 2. All data should be entered into the TEAM database as soon as possible after collection. The fields in the database are often constrained to a pulldown menu of only a given set of available choices. For example, The following fields will be included in the database, along with the possible choices and their definitions. The types of data fields that appear on the forms is of four types: transect information, ecological information, specimen information, and species information. The data forms in the database are divided somewhat differently, but the fields that appear are listed. Information about the transect and its location FIELD CHOICES DEFINITION Array ID

1-6 Identity of the array

Transect Coordinates Identifies the grid square where trap is located. One set of coordinates identifies the square because the second set of coordinates is assumed to be the next higher number of each.

Sampling type Winkler Pitfall

Type of sampling

Section of transect 1-10 40-50 90-100

Section of transect where ecological data was taken


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Information about ecological conditions at time of sampling. (This will also include all the data listed on field data sheet in Appendix 2.) FIELD CHOICES DEFINITION Date dd/mm/yr Date of sampling Time of day hh:mm Time at start of sampling Names of collectors Name1

Name 2 Names of field workers

Air temperature XX.X° C Air temperature at onset of transect sampling

Relative humidity XX.X % Relative humidity at onset of transect sampling

# stems < 2 m height (etc for ecological data)

Count data Number of stems counted in this category

Information about specimens sampled Genus and species (list of possible species, including

morphospecies A,B, etc) Identity of each specimen collected. If a species is encountered that does not appear on the list, you must first enter the species’ information in the species table below. It will then automatically appear in the pulldown list.

Sex M F undetermined

Sex of specimen, if known

Date determined dd/mm/yr Date of the identification of the specimen

Determined by Name 1 etc

Name of person who identified the specimen

Information about species FIELD CHOICES DEFINITION Order Hymenoptera Only one order sampled

Family Formicidae Formicidae only is the target group.

Sub-family list Sub-family

Genus and species Each species, even morphospecies, needs a new entry before being able to record its specimens

Species authority Name of the author who first described the species

There are many more fields for species specific information that is necessary for taxonomists and museum collections, but for the purposes of this protocol, what is listed above is sufficient.


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Data analysis See Avian Protocol for a discussion of analytical methods of species richness estimators. Further analyses will include ordination methods of diversity indices along axes that describe climate, soil, and vegetation measures; examination of differences in community composition between years, and between regions using rarefaction methods (Gotelli and Graves 1996), similarity indices, and ordination (Kremen 1992; Kremen 1994). More in-depth explanations of data analyses will be given in future versions of this document.

Acknowledgements Discussions with Leeanne Alonso, Roberto Brandão, Brian Fisher, Ana Harada, and Jack Longino have contributed to the ideas and methods described. Field tests with Ana Harada and Bill Overal helped to fine tune many of important details of the protocol.


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Literature Cited Agosti, D., J. D. Majer, et al. (2000). Ants: Standard Methods for Measuring and

Monitoring Biodiversity, Smithsonian Institution Press. Alonso, L. (2000). Ants as Indicators of Diversity. Ants: Standard Methods for

Measuring and Monitoring Biodiversity. D. Agosti, J. D. Majer, L. E. Alonso and T. R. Schultz. Washington and London, Smithsonian Institution Press.

Bandeira, A. and A. Y. Harada (1998). "Densidade e distribuição vertical de macroinvertebrados em solos argilosos e arenosos na Amazônia central." Acta Amazonica 28(2): 191-204.

Bazzaz, F. A. (1998). "Tropical forests in a future climate: Changes in biological diversity and impact on the global carbon cycle." Climatic Change 39(2-3): 317-336.

Bestelmeyer, B. T., D. Agosti, et al. (2000). Field techniques for the study of ground dwelling ants: an overview, description, and evaluation. Ants: Standard Methods for Measuring and Monitoring Biodiversity. D. Agosti, J. D. Majer, L. E. Alonso and T. R. Schultz. Washington and London, Smithsonian Institution Press.

Bestelmeyer, B. T. and J. A. Wiens (1996). "The effects of land use on the structure of ground-foraging ant communities in the Argentine Chaco." Ecological Applications 6(4): 1225-1240.

Bestelmeyer, B. T. and J. A. Wiens (2001). "Ant biodiversity in semiarid landscape mosaics: The consequences of grazing vs. natural heterogeneity." Ecological Applications. 11(4): 1123-1140.

Catangui, M. A., B. W. Fuller, et al. (1996). "Abundance, diversity, and spatial distribution of ants (Hymenoptera: Formicidae) on mixed-grass rangelands treated with diflubenzuron." Environmental Entomology 25(4): 757-766.

Delabie, J. H. C., B. L. Fisher, et al. (2000). Sampling effort and choice of methods. Ants: Standard Methods for Measuring and Monitoring Biodiversity. D. Agosti, J. D. Majer, L. E. Alonso and T. R. Schultz. Washington and London, Smithsonian Institution Press.

Enquist, C. A. F. (2002). "Predicted regional impacts of climate change on the geographical distribution and diversity of tropical forests in Costa Rica." Journal of Biogeography 29(4): 519-534.

Erwin, T. (1989). "Sorting tropical forest canopy samples." Insect Collection News 2(1): 8.

Feener, D. and E. Schupp (1998). "Effect of treefall gaps on the patchiness and species richness of Neotropical ant assemblages." Oecologia 116(1-2): 191-201.

Fittkau, E. J. and H. Klinge (1973). "On biomass and trophic structure of the central Amazonian rain forest ecosystem." Biotropica 5(1): 2-14.

Gotelli, N. J. and G. G. Graves (1996). Null models in ecology. Washington, Smithsonian Institution Press.

Holldobler, B. and E. O. Wilson (1990). The Ants. Cambridge MA, Belknap Press of Harvarad University Press.

Kaspari, M., L. Alonso, et al. (2000). "Three energy variables predict ant abundance at a geographical scale." Proceedings of the Royal Society of London, Series B: Biological Sciences. 267(1442): 485-489.


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Kaspari, M., S. O'Donnell, et al. (2000). "Energy, Density, and Constraints to Species Richness: Ant Assemblages along a Productivity Gradient." American Naturalist. 155(2): 280-293.

Kaspari, M. and M. D. Weiser (2000). "Ant Activity along Moisture Gradients in a Neotropical Forest." Biotropica 32(4a): 703-711.

Kremen, C. (1992). "Assessing the Indicator Properties of Species Assemblages for Natural Areas Monitoring." Ecological Applications 2(2): 203-217.

Kremen, C. (1994). "Biological Inventory Using Target Taxa: A Case Study of the Butterflies of Madagascar." Ecological Applications 4(3): 407-422.

Majer, J. D. and J. H. C. Delabie (1994). "Comparison of the ant communities of annually inundated and terra firme forests at Trombetas in the Brazilian Amazon." Insectes Sociaux 41: 343-359.

Majer, J. D., J. H. C. Delabie, et al. (1997). "Ant litter fauna of forest, forest edges and adjacent grassland in the Atlantic rain forest region of Bahia, Brazil." Insectes Sociaux 44(3): 255-266.

Morrison, L. (1998). "The spatiotemporal dynamics of insular ant metapopulations." Ecology 79(4): 1135-1146.

Nepstad, D. C. (1995). Forest recovery following pasture abandonment in Amazonia: Canopy seasonality, fire resistance, and ants. Evaluating and monitoring the health of large-scale ecosystems. C. L. Rapport, C. L. Gaudet and P. Calow. Berlin and Heidelberg, Springer-Verlag.

Parmesan, C. and G. Yohe (2003). "A globally coherent fingerprint of climate change impacts across natural systems." Nature 421(6918): 37-42.

Root, T. L. and S. H. Schneider (2002). Climate Change: Overview and Implications for Wildlife. Wildlife Responses to Climate Change: North American Case Studies. S. H. Schneider and T. L. Root. Washington, Island Press.

Walther, G. R., E. Post, et al. (2002). "Ecological responses to recent climate change." Nature 416(6879): 389-395.


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Appendix I Equipment list and Suppliers in U.S.

For litter sampling (per transect): random number table sighting compass 100 m nylon rope knotted at 10 m intervals blue flagging 1 1m2 quadrat made of wood or PVC (movable joints; one corner should be openable) 1 machete 2 pair gloves 1 small hand rake 1 small hand trowel 1 watch with seconds or stopwatch 1 litter sifter 10 litter stuff sacks (included with the mini-Winkler apparatus) and some extras 2 waterproof notebooks pencils large plastic bag, large enough to carry 10 mini-Winkler samples without compressing them For pitfall traps (per transect): soil auger (or sharp edged pipe with diameter of the pitfall cups) to dig pitfall trap holes; must be strong enough to cut through roots. 10 durable plastic cups, 7-8 cm diameter, 8-10 cm depth, with no ridges inside or at mouth 1 liter 75% Ethanol 5 ml liquid detergent 10 small plastic plates, few cm larger than diameter of cups; used as a roof for the pitfall trap thin wooden skewers labels (card stock) squeeze bottle forceps waterproof and Ethanol proof ink pen For measuring ecological conditions Kestrel 3000 Pocket Weather Station Electronic soil probes for pH, temperature, humidity Small ruler (15 cm) For processing mini-Winkler litter samples and pitfall trap contents (per transect): 2 pieces nylon rope (5 m) 1 2x3 m plastic sheet (only 2 are needed in total) 10 mini-Winkler apparatus (each mini-Winkler contains a mesh bag, and the external cloth portion) and a few extra 2 light colored plastic trays medium width (5 cm) paintbrush fine (2-3 mm) paintbrush forceps 10 plastic cups with no ridges inside or at mouth 2 liters 95% EtOH (source alcohol, not denatured EtOH) squirt bottle 10 vials (25-50 ml) with tight caps 1 large ziploc bag labels (card stock) waterproof and EtOH proof ink pen scissors


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Many of the items on the list are readily available in most cities. Some catalogue or web order suppliers for items which may be difficult to find in some areas are listed below.

For mini-Winkler apparatus:

Marizete Pereira dos Santos Km 22 rodovia Ilhéus-Itabuna cx Postal-07 Cep-45600-000 Ministério da Agricultura - CEPLAC Centro de Pesquisa do Cacau SECEN Residence address: Rua do Coqueiro nº60 Bairro- Conquista Ilhéus - Bahia Cep - 45 650-000 Email: "Paulo Roberto Pires Santos" <[email protected]>

For waterproof pens, waterproof notebooks, specimen processing and identification tools, and any other general entomological equipment:

BioQuip Products 2321 Gladwick Street Rancho Dominguez, CA 90220 USA Telephone: 310-667-8800 Fax: 310-667-8808 e-mail: [email protected] web: www.bioquip.com

For compass, flagging, waterproof notebooks, pocket weather station, and other general field supplies:

Forestry Suppliers, Inc. Post Office Box 8397 Jackson Mississippi 39284-8397 USA Telephone (USA): 800-647-5368 Fax (USA): 800-543-4203 Telephone (International): 601-354-3565 e-mail for international customers: [email protected] Web: www.forestry-suppliers.com Ben Meadows Company

PO Box 5277 Janesville WI 53547-5277 USA Telephone (USA & Canada): 800-241-6401 Telephone (Worldwide): 608-743-8001 Fax (USA & Canada): 800-628-2068 Fax (Worldwide): 608-743-8007 E-mail for international customers: [email protected] Web: www.benmeadows.com


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Appendix II Field data form for leaf litter transects

DATE: (date-Month-year) NAMES OF COLLECTORS: ARRAY #: COORDINATES OF TRANSECT: STARTING TIME: (hh:mm) AIR TEMPERATURE AT STARTING TIME: RELATIVE HUMIDITY AT STARTING TIME: VEGETATION DESCRIPTION (NUMBER OF LIVE STEMS) INTERVAL 1-10 m INTERVAL 40-50 m INTERVAL 90-100 m <1 m HEIGHT

< 2 m HEIGHT

>2 m HEIGHT

>10 cm DBH

CANOPY DESCRIPTION INTERVAL 1-10 m INTERVAL 40-50 m INTERVAL 90-100 m APPROXIMATE HEIGHT (m)

APPROXIMATE HEIGHT OF CANOPY EMERGENTS (m)

AMOUNT OF COVER (CLOSED, PARTIALLY OPEN, OPEN GAP, ETC)

HABITAT DESCRIPTION (% GROUND COVER OF EACH TYPE) INTERVAL 1-10 m INTERVAL 40-50 m INTERVAL 90-100 m SOIL

LIVE VEGETATION INCLUDING ROOTS

DECOMPOSING WOOD

STONE

LEAF LITTER

LEAF LITTER CONDITIONS INTERVAL 1-10 m INTERVAL 40-50 m INTERVAL 90-100 m pH TEMPERATURE RELATIVE HUMIDITY DEPTH (cm)


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Figures

Figure 1. Placement of ant and termite transects in 1 km2 array. Dotted lines indicate the divisions among four quadrants of the IMA, within each of which will be a randomly placed. transect Blue lines indicate transects which run parallel to each other 3 meters apart.


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Figure 2. 1m2 quadrat for leaf litter ant sampling. Litter has been collected in center of square.

Figure 3. Sifting litter (left), and fine litter after having been sifted (right). The litter sample will be further processed in the lab to extract the ants from it.


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Figure 4. Pitfall trap with mouth flush with the ground (left), and roof of pitfall trap made of plastic plate and wooden skewers.


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Figure 5. Transferring litter samples into mini-Winkler mesh sack. Care must be taken to transfer all the contents of the litter sample, using a paintbrush to sweep fine particles that fall through back into the mesh.


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Figure 6. Mini-Winkler extractors (upper). The mesh sack contains the litter sample, and remains relatively flat as it hangs vertically, centered inside the mini-Winkler. (Lower left photograph is a view from the top down into the mini-Winkler). The bottom of the mini-Winkler holds a small cup with a sample label and alcohol (lower right), into which the ants fall when they exit the mesh sack.

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