The author(s) thank the University of Wisconsin Biotechnology Center DNA Sequencing Facility for providing amplification and sequencing facilities and services, Caitlin Carlson, Jennifer Knack, Jake Otto, and Max Haase for providing technical assistance with molecular analysis. This work was supported by USDA-Agricultural Research Service appropriated funds (Current Research Information System #3655-21220-001). Further support was provided by the National Science Foundation (under Grant No. DEB-1442148), the DOE Great Lakes Bioenergy Research Center (DOE Office of Science BER DE-FC02-07ER64494), and the USDA National Institute of Food and Agriculture (Hatch project 1003258). C.T.H. is a Pew Scholar in the Biomedical Sciences and an Alfred Toepfer Faculty Fellow, supported by the Pew Charitable Trusts and the Alexander von Humboldt Foundation, respectively. The authors thank Hannah Gaines-Day and Olivia Bernauer for their contributions to our earlier, published field work.

1. Examine the Effect of Fungicide Exposure on Bumble Bee Colony Success Using Field Cage Experiments
<ol>
	<li>Set up ten mesh cages in a field planted with oats. Dig a trench around each cage, and dig all four edges of the mesh cage into the ground to ensure that bees cannot escape. Stock the cages with potted, flowering plants that are known to be attractive to bees (e.g. buckwheat, borage, alyssum, cosmos, and sunflowers) (Figure 2).</li>
	<li>Supplement the cages with a single tray (36 cm x 42 cm) of in-bloom clover. Cluster floral resources within one corner of the cage, occupying a space approximately 2.5 m x 1 m. Vegetate the remaining cage area by the oats.</li>
	<li>Randomly assign the bumble bee colonies, each containing workers and a single queen, a treatment (fungicide present/absent, N = 5 colonies per treatment, 10 colonies total), then place them within a field cage (N = 1/cage) for 29 days (23 June - 21 July 2014).</li>
	<li>Orient the colony boxes such that the colony&#39;s openings point to the south to provide the bees with optimal navigational conditions. Subsidize the colonies with sugar water bladders, placed inside the hive boxes to supplement nectar availability.</li>
	<li>Apply chlorothalonil-based fungicide at a field-relevant level (20 g/L) to flowering plants in the five fungicide treatment cages, using a hand held pesticide sprayer, twice during the study (day 0 and 13). Coat the flowers uniformly such that no further liquid adheres to floral surfaces (Figure 3).</li>
	<li>At the conclusion of the field cage study, remove the B. impatiens colonies from the cages by hand, cool the hives by placing in -20 &#176;C freezer for 20 min.</li>
	<li>Remove bees using sterile forceps and record the number of larvae, pupae, adult females (i.e. foragers), and adult males. Using an analytical balance record the dry-weight of the mother queen, larvae, pupae, adult females (i.e. foragers), and adult males.</li>
</ol>
2. Examine the Effects of Fungicide Exposure on Microbial Communities in the Pollen-provisions of Bumble Bee Nests Using Laboratory Based In Vivo Trials
<ol>
	<li>Pulverize commercially purchased pollen to a fine powder using a standard laboratory ball-mill. Sterilize powdered pollen by soaking in 70% ethanol, allowing it to evaporate overnight under UV light. Verify sterility of pollen by plating ~0.5 mg on general-purpose agar media.
	<ol>
		<li>To the dry, sterilized pollen, using sterilized pipettes, add field relevant dosage of fungicide: propiconazole at 14.3%; azoxystrobin at 22.9% for treatments (0.74 &#181;L and 0.65 &#181;L respectively/ day/ hive). Mix well using sterilized wooden sticks.</li>
	</ol>
	</li>
	<li>Place 6 experimental hives (n = 3 each for control and treatment) in a clean, hygienic laboratory benchtop maintained at room temperature. Each day weigh 4.27 g of pollen34,35 mixed with fungicides (for treatments) or sterile water (for control) inside a hood using standard aseptic technique.
	<ol>
		<li>Using the trap doors provided by the side of the cardboard box enclosing the hives, introduce the pollen inside the hives. Supplement the hives with sterilized sugar solution each week. Continue feeding regime for four weeks.</li>
	</ol>
	</li>
	<li>At the conclusion of the lab-based study, cool the hives by placing in -20 &#176;C freezer for 20 min. Scrape out the pollen-provisions contained within brood chambers using sterilized forceps and spatulas and place in sterile storage tubes. Store at -80 &#176;C. Count and record the weight of the workers and queen mother at the start and end of the experiment (step 1.7).</li>
	<li>Isolate DNA from pollen-provision sample using commercially available DNA isolation kits (see Materials Table for details).
	<ol>
		<li>Add 0.25 g of pollen-provision to extraction tubes, briefly vortex to mix.</li>
		<li>Make a 200 mg/mL lysozyme solution in deionized distilled water, enough for 50 &#181;L/sample. Shake vigorously to completely form solution.</li>
		<li>Add 50 &#181;L of the lysozyme solution to the extraction tubes with sample in it, and mix well by inverting several times. Incubate the tube for 10 min at 37 &#176;C in a water bath. If precipitate has formed, heat solution to 60 &#176;C until dissolved before use.</li>
		<li>Add 70 &#181;L of aqueous lysis solution to extraction tubes, secure horizontally on a flatbed vortex pad with tape, and vortex for 10 min. Centrifuge tubes at 10,000 x g for 30 s at room temperature.</li>
		<li>Transfer the supernatant to a clean 2 mL collection tube. Add 250 &#181;L of protein precipitation solution, and vortex for 5 s. Incubate at 4 &#176;C for 5 min in an ice bath. 
		NOTE: Expect between 400 &#181;L to 500 &#181;L of supernatant. Supernatant may still contain some particles.</li>
		<li>Centrifuge the tubes at room temperature for 1 min at 10,000 x g, and transfer up to 600 &#181;L of supernatant to a clean 2 mL collection tube. Add 200 &#181;L of aqueous inhibitor removal solution, vortex briefly, and incubate at 4 &#176;C for 5 min. Centrifuge the tubes at room temperature for 1 min at 10,000 x g. Transfer up to 750 &#181;L of supernatant into a clean 2 mL collection tube.</li>
		<li>Add 1200 &#181;L of aqueous bind solution to the supernatant, and vortex for 5 s. Load approximately 675 &#181;L of the supernatant onto a spin filter, and centrifuge at 10,000 x g for 1 min at room temperature. Discard the flow through.</li>
		<li>Repeat 2.4.7. twice. 
		NOTE: A total of three loads for each sample processed are required.</li>
		<li>Add 500 &#181;L ethanol, and centrifuge at room temperature for 30 s at 10,000 x g. Discard the flow through, and centrifuge again at room temperature for 1 min at 10,000 x g. Place spin filter in a clean 2 mL collection tube.</li>
		<li>Add 100 &#181;L of elution buffer to the center of the filter membrane. Centrifuge at room temperature for 30 s at 10,000 x g. Discard the spin filter. Store collected DNA between -20 &#176;C and -80 &#176;C</li>
	</ol>
	</li>
	<li>Use isolated DNA for sequencing.
	<ol>
		<li>Quantify, and normalize isolated DNA to 2 ng/&#181;L by fluorometric analysis.
		<ol>
			<li>Prepare reactions in triplicate for each extracted sample to compare the relative amounts of 28S (plant), analyze ITS (fungal), and 16S (bacterial) components of each pollen-provision sample36. Ensure that each reaction contains 10 ng of total DNA, 2x of asymmetrical cyanine dye based master mix, and 2.5 &#181;L each of forward and reverse primers pairs 28KJ/28B37 for plant and ITS1/ITS5.8R for fungal DNA38.</li>
		</ol>
		</li>
		<li>Amplify DNA using the following parameters: 2 min pre-denaturation at 50 &#176;C, 2 min initial denaturation at 95 &#176;C, 40 cycles of (15 s at 98 &#176;C, 15 s at 58 &#176;C, 60 s at 72 &#176;C), followed by a melt curve.</li>
		<li>Prepare a 2-step, nested PCR protocol using next-generation sequencing libraries targeting the 16S rRNA V3/V4 variable region and ITS/5.8s rRNA spacer region.
		<ol>
			<li>Add 12.5 &#181;L of DNA to 5 pmol of each primer and 2x master mix. Make separate reactions for each region (16S or ITS; see Table 1). Modify region specific primers as previously described39,40 to add sequencer-specific adapter overhang nucleotide sequences to the gene-specific sequences.</li>
			<li>Perform initial amplification using the following parameters: 3 min initial denaturation at 95 &#176;C, 25 cycles of (30 s at 95 &#176;C, 30 s at 55 &#176;C, 30 s at 72 &#176;C), 5 min final extension at 72 &#176;C.</li>
			<li>Following initial amplification, verify library size, and quantity by electrophoretic mobility, and clean using a 1x volume solid phase reversible immobilization beads to remove residual primers and reaction reagents. Pool 16S and ITS amplicons quantitatively to create a single amplicon pool for each sample.</li>
			<li>Add sequencer specific adapters and sample specific dual indexes using the following primers (see Table 1). Add 2.5 &#181;L of amplified DNA to 5 pmol of each primer and 2x master mix. Perform library amplification using the following parameters: 3 min initial denaturation at 95 &#176;C, 8 cycles of (30 s at 95 &#176;C, 30 s at 55 &#176;C, 30 s at 72 &#176;C), 5 min final extension at 72 &#176;C.</li>
		</ol>
		</li>
		<li>Following PCR, clean finished libraries using a 1 x volume of solid phase reversible immobilization beads. Assess quality and quantity of the finished libraries using electrophoretic mobility, and fluorometry, respectively. Standardize libraries to 2 &#181;M and pool prior to sequencing.</li>
		<li>Perform next-generation sequencing on a platform analogous to the adapters added in the secondary PCR, with an appropriate length to cover the entire amplicon41.</li>
	</ol>
	</li>
	<li>Sequence annotation and microbial composition analysis.
	<ol>
		<li>Combine pair-end sequencing data (R1 and R2) into single contigs for all sequenced libraries. After merging both R1 and R2 files, a single fasta file for each of the libraries is produced. 
		NOTE: This step and the processes described below (unless otherwise indicated) are performed in Mothur version 1.38.038.</li>
		<li>Screen each fasta file to remove ambiguous bases, unexpected long sequences, and long homopolymers. 
		NOTE: The parameters for screening, and sequence removal are maxambig = 0, maxlength = 600, and maxhomop = 8 for both genes. Remove identical sequences, but keep a contingence table separately for all the libraries (a.k.a. count table in Mothur).</li>
		<li>Align unique sequences of the gene 16S library against the SILVA database version 123, and the gene ITS library against the UNITE ITS database42. 
		NOTE: Sequences must be annotated from the kingdom level to the genus level.
		<ol>
			<li>Subsequently, cluster aligned sequences with the function pre.cluster using diff=5, and remove chimeras.</li>
		</ol>
		</li>
		<li>Classify sequences using the method Wang43 based on the SILVA (for gene 16S) and UNITE ITS (for gene ITS) taxonomy files (cutoff value of 80%).</li>
		<li>Load into R44 the contingence tables (one per gene) generated during the classification process in Mothur. At each taxonomic level, obtain relative abundance per library for further analysis of microbial community changes. 
		NOTE: At each taxonomic level, merge together taxonomic groups when relative abundance is less than 2% for all the experimental repetitions.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

Investigations into the effects of fungicides on bee health have remained an understudied aspect of pest management strategies. Our study aims to bridge this knowledge gap by using a suite of complementary techniques that explicitly isolate the potential factors driving bee declines. The planning, rationale, and rendering of these experiments are detailed below.
It is important to ensure that no bees are allowed to escape the mesh of the cage experiments, since this would compromise demography...

Managed and wild bee species are experiencing widespread declines, with major implications for both natural and agricultural systems1. Despite concerted efforts to understand the causes of this problem, the factors driving honey bee declines are still not well understood2,3,4. For certain species of wild, native bees, the situation has become dire5,6. If bee populations cannot be sustained when they intersect with industrial agriculture, their populations will continue to fall, and the crops requiring pollinators (35% of worldwide production7) will endure reduced harvests.
While many potential factors such as pesticide exposure, disease, and habitat loss1,4,8,9,10 have been implicated in honey bee decline, relatively little is known about the interactive effect of these stressors on native bees health, within or near agricultural systems. Many current research efforts continue to focus on insecticides, (e.g., neonicotinoids11,12), although past research indicates that fungicides may also play a role in bee decline by impairing memory formation, olfactory reception13, nest recognition14, enzyme activity and metabolic functions15,16,17. Globally, fungicides continue to be applied to flowering crops during bloom. Recent studies have documented that bees commonly bring fungicide residues back to the hive18, indeed, studies have shown a large proportion of tested hives contained fungicide residues19,20. Further work has revealed that fungicide residue is associated with high rates of honey bee larval mortality21,22,23 and the presence of &#34;entombed pollen&#34; within colonies, which although non-toxic, is devoid of microbial activity and is nutritionally compromised24. Despite the fact that fungicides have long been considered &#34;bee-safe,&#34; there is now evidence that exposure to fungicide alone can cause severe colony losses in a native bumble bee species, Bombus impatiens25.
To establish causality between fungicide exposure and colony mortality, the modus operandi of these chemicals need to be determined. As evidenced in soils26, sediments27, and aquatic environments28, by targeting fungi, fungicides most likely alter fungal abundance and diversity within pollen-provisions, thereby invoking a major community shift that may strongly favor bacteria. Without fungal competitors or antagonists, certain pathogenic bacteria can proliferate relatively unchecked, facilitating the spoilage of pollen-provisions. Past research has demonstrated that microorganisms, particularly yeasts and filamentous fungi, serve as nutritional symbionts for bees29,30,31, protect against parasites and pathogens32,33, and provide long-term preservation of pollen stores. Fungicides, therefore, may indirectly harm immature bees by disrupting the microbial community that is needed to provide these services and/or by increasing susceptibility to opportunistic pathogens and parasites12. With increasing demands on food production, crops worldwide are being sprayed each year with fungicides during bloom, underscoring the need to understand the magnitude of such fungicide-induced effects.
To-date, the primary knowledge gaps relating to native bee microbial ecology can be represented by the following questions: To what extent does fungicide change the microbial community within bee pollen-provisions? What are the downstream impacts of consuming pollen with a profoundly altered microbial community? In keeping with these ecologically germane questions, experiments were developed with the primary goals of revealing 1) that fungicide residue alone can cause severe colony decline in a native bee species; 2) the degree to which microbial communities in pollen-provisions are altered by fungicides, and 3) how bee health is affected by a severely altered microbial community. The experimental objectives were defined to address the above questions using a combination of laboratory- and field-based experiments. Using state-of-the-art metagenomic and molecular techniques alongside traditional methods of field observation, this research aims to piece together the potential effects of fungicides on bee health.
The first objective of this study is to demonstrate that fungicide exposure alone can cause significant colony losses among native bee species. A study involving large field cages was used to investigate the effects of fungicide exposure on the colony growth of Bombus impatiens, a ubiquitous, abundant native bee in the US (Figure 1, Figure 2, Figure 3). It was hypothesized that fungicide-treated hives would present lower fitness, and atypical demography compared to non-exposed hives. Data obtained from this experiment supported this hypothesis, demonstrating that fungicide residues within pollen can be the sole cause of profound colony losses in a native bumble bee species25. The second objective of this study is to investigate the response of the pollen microbiome to fungicide exposure. It is hypothesized that the community composition of microbes within pollen-provisions exposed to fungicides will be different from that of untreated pollen. While fungal abundance and diversity are expected to decline significantly, bacteria and/or a single dominant fungal species will likely grow unchecked in the absence of other competing fungi. Through a series of in-vivo trials, these shifts in microbial community composition will be analyzed using metagenomics.

<table>
	<tbody>
		<tr>
			<td>Natupol Beehive</td>
			<td>Koppert</td>
			<td>USRESM1</td>
			<td>16 hives</td>
		</tr>
		<tr>
			<td>Propiconazole 14.3</td>
			<td>Quali-Ppro</td>
			<td>60207-90-1</td>
			<td>Propiconazole 14.3%</td>
		</tr>
		<tr>
			<td>Abound</td>
			<td>Syngenta</td>
			<td>4033540</td>
			<td>Azoxystrobin 22.9%</td>
		</tr>
		<tr>
			<td>Chlorothalonil</td>
			<td>Syngenta</td>
			<td>3452</td>
			<td>Fungicide used for trials</td>
		</tr>
		<tr>
			<td>Pollen granules</td>
			<td>Bee rescued</td>
			<td>B004D5650C</td>
			<td>3X 16oz bottles, pollen for trials</td>
		</tr>
		<tr>
			<td>Bacterial strains for inoculation</td>
			<td>Currie Lab</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Yeast strains for inoculation</td>
			<td>Hittinger lab</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Primer pairs</td>
			<td>UW Biotech Center</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DNA Isolation Kit</td>
			<td>Mo Bio</td>
			<td>12830-50</td>
			<td>Commercial DNA isolation kit</td>
		</tr>
		<tr>
			<td>Qubit dsDNA HS Assay Kit</td>
			<td>Thermo Fisher</td>
			<td>Q32851</td>
			<td>DNA quantification tool</td>
		</tr>
		<tr>
			<td>Select Master Mix for CFX</td>
			<td>Thermo Fisher</td>
			<td>4472952</td>
			<td>Used to perform real-time PCR using SYBR GreenER dye.</td>
		</tr>
		<tr>
			<td>Real-Time PCR Detection System</td>
			<td>Bio Rad</td>
			<td>1855196</td>
			<td>Instrument used for PCR amplification</td>
		</tr>
		<tr>
			<td>PCR Clean-Up Kit,</td>
			<td>Axygen</td>
			<td>10159-696</td>
			<td>Used for efficient removal of unincorporated dNTPs, salts and enzymes</td>
		</tr>
		<tr>
			<td>DNA 1000 Kit</td>
			<td>Agilent</td>
			<td>5067-1504</td>
			<td>Used for sizing and analysis of DNA fragments</td>
		</tr>
		<tr>
			<td>MiSeq Sequencer</td>
			<td>Illumina</td>
			<td></td>
			<td>Used for next-generation sequencing</td>
		</tr>
		<tr>
			<td>Assorted glassware (beaker, flasks, pipettes, test tubes, repietters)</td>
			<td>VWR</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

empirical, metagenomic, and computational techniques illuminate the mechanisms by which fungicides compromise bee health

Growers often use fungicide sprays during bloom to protect crops against disease, which exposes bees to fungicide residues. Although considered &#34;bee-safe,&#34; there is mounting evidence that fungicide residues in pollen are associated with bee declines (for both honey and bumble bee species). While the mechanisms remain relatively unknown, researchers have speculated that bee-microbe symbioses are involved. Microbes play a pivotal role in the preservation and/or processing of pollen, which serves as nutrition for larval bees. By altering the microbial community, it is likely that fungicides disrupt these microbe-mediated services, and thereby compromise bee health. This manuscript describes the protocols used to investigate the indirect mechanism(s) by which fungicides may be causing colony decline. Cage experiments exposing bees to fungicide-treated flowers have already provided the first evidence that fungicides cause profound colony losses in a native bumble bee (Bombus impatiens). Using field-relevant doses of fungicides, a series of experiments have been developed to provide a finer description of microbial community dynamics of fungicide-exposed pollen. Shifts in the structural composition of fungal and bacterial assemblages within the pollen microbiome are investigated by next-generation sequencing and metagenomic analysis. Experiments developed herein have been designed to provide a mechanistic understanding of how fungicides affect the microbiome of pollen-provisions. Ultimately, these findings should shed light on the indirect pathway through which fungicides may be causing colony declines.

Field cage study:
Data obtained from the cage experiments showed that the bumble bee colonies had a significant response to fungicide exposure. The fungicide-treated hives produced significantly fewer workers (12.2 &#177; 3.8, mean &#177; SE) than the control hives (43.2 &#177; 11.2, F1,9= 6.8, p = 0.03) (Figure 4). Additionally, the bee biomass of the fungicide-tr...

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Empirical, Metagenomic and Computational Techniques Illuminate the Mechanisms by which Fungicides Compromise Bee Health | Environment | Video

Empirical, Metagenomic, and Computational Techniques Illuminate the Mechanisms by which Fungicides Compromise Bee Health

Microbial consortia within bumble bee hives enrich and preserve pollen for bee larvae. Using next generation sequencing, along with laboratory and field-based experiments, this manuscript describes protocols used to test the hypothesis that fungicide residues alter the pollen microbiome, and colony demographics, ultimately leading to colony loss.

Growers often use fungicide sprays during bloom to protect crops against disease, which exposes bees to fungicide residues. Although considered ...

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9.6K Views.  USDA-ARS. Microbial consortia within bumble bee hives enrich and preserve pollen for bee larvae. Using next generation sequencing, along with laboratory and field-based experiments, this manuscript describes protocols used to test the hypothesis that fungicide residues alter the pollen microbiome, and colony demographics, ultimately leading to colony loss.

Video: Empirical, Metagenomic, and Computational Techniques Illuminate the Mechanisms by which Fungicides Compromise Bee Health

Optimized Negative Staining: a High-throughput Protocol for Examining Small and Asymmetric Protein Structure by Electron Microscopy

Structural determination of proteins is rather challenging for proteins with molecular masses between 40 - 200 kDa. Considering that more than half of natural proteins have a molecular mass between 40 - 200 kDa1,2, a robust and high-throughput method with a nanometer resolution capability is needed. Negative staining (NS) electron microscopy (EM) is an easy, rapid, and qualitative approach which has frequently been used in research laboratories to examine protein structure and protein-protein interactions. Unfortunately, conventional NS protocols often generate structural artifacts on proteins, especially with lipoproteins that usually form presenting rouleaux artifacts. By using images of lipoproteins from cryo-electron microscopy (cryo-EM) as a standard, the key parameters in NS specimen preparation conditions were recently screened and reported as the optimized NS protocol (OpNS), a modified conventional NS protocol 3 . Artifacts like rouleaux can be greatly limited by OpNS, additionally providing high contrast along with reasonably high&#8208;resolution (near 1 nm) images of small and asymmetric proteins. These high-resolution and high contrast images are even favorable for an individual protein (a single object, no average) 3D reconstruction, such as a 160 kDa antibody, through the method of electron tomography4,5. Moreover, OpNS can be a high&#8208;throughput tool to examine hundreds of samples of small proteins. For example, the previously published mechanism of 53 kDa cholesteryl ester transfer protein (CETP) involved the screening and imaging of hundreds of samples 6. Considering cryo-EM rarely successfully images proteins less than 200 kDa has yet to publish any study involving screening over one hundred sample conditions, it is fair to call OpNS a high-throughput method for studying small proteins. Hopefully the OpNS protocol presented here can be a useful tool to push the boundaries of EM and accelerate EM studies into small protein structure, dynamics and mechanisms.

More than half of proteins are small proteins (molecular mass &#60;200 kDa) that are challenging for both electron microscope imaging and three-dimensional reconstructions. Optimized negative staining is a robust and high-throughput protocol to obtain high contrast and relatively high resolution (~1 nm) images of small asymmetric proteins or complexes under different physiological conditions.

Structural determination of proteins is rather challenging for proteins with molecular masses between 40 - 200 kDa. Considering that more than half of ...

Spotting Cheetahs: Identifying Individuals by Their Footprints

The cheetah (Acinonyx jubatus) is Africa's most endangered large felid and listed as Vulnerable with a declining population trend by the IUCN1. It ranges widely over sub-Saharan Africa and in parts of the Middle East. Cheetah conservationists face two major challenges, conflict with landowners over the killing of domestic livestock, and concern over range contraction. Understanding of the latter remains particularly poor2. Namibia is believed to support the largest number of cheetahs of any range country, around 30%, but estimates range from 2,9053 to 13,5204. The disparity is likely a result of the different techniques used in monitoring.
Current techniques, including invasive tagging with VHF or satellite/GPS collars, can be costly and unreliable. The footprint identification technique5 is a new tool accessible to both field scientists and also citizens with smartphones, who could potentially augment data collection. The footprint identification technique analyzes digital images of footprints captured according to a standardized protocol. Images are optimized and measured in data visualization software. Measurements of distances, angles, and areas of the footprint images are analyzed using a robust cross-validated pairwise discriminant analysis based on a customized model. The final output is in the form of a Ward's cluster dendrogram. A user-friendly graphic user interface (GUI) allows the user immediate access and clear interpretation of classification results.
The footprint identification technique algorithms are species specific because each species has a unique anatomy. The technique runs in a data visualization software, using its own scripting language (jsl) that can be customized for the footprint anatomy of any species. An initial classification algorithm is built from a training database of footprints from that species, collected from individuals of known identity. An algorithm derived from a cheetah of known identity is then able to classify free-ranging cheetahs of unknown identity. The footprint identification technique predicts individual cheetah identity with an accuracy of &gt;90%.

The cheetah (Acinonyx jubatus) is an iconic, endangered species, but conservation efforts are challenged by habitat shrinkage and conflict with commercial farmers. The footprint identification technique, a robust, accurate and cost-effective image classification system, is a new approach to monitoring cheetahs.

The cheetah (Acinonyx jubatus) is Africa's most endangered large felid and listed as Vulnerable with a declining population trend by the IUCN1. It ranges ...

Evaluating the Effect of Environmental Chemicals on Honey Bee Development from the Individual to Colony Level

The presence of pesticides in the beekeeping environment is one of the most serious problems that impacts the life of a honey bee. Pesticides can be brought back to the beehive after the bees have foraged on flowers that have been sprayed with pesticides. Pesticide contaminated food can be exchanged between workers which then feed larvae and therefore can potentially affect the development of honey bees. Thus, residual pesticides in the environment can become a chronic damaging factor to honey bee populations and gradually lead to colony collapse. In the presented protocol, honey bee feeding methods are described and applied to either an individual honey bee or to a colony. Here, the insect growth regulator (IGR) pyriproxyfen (PPN), which is widely used to control pest insects and is harmful to the development of honey bee larvae and pupae, is used as the pesticide. The presenting procedure can be applied to other potentially harmful chemicals or honeybee pathogens for further studies.

Herein we present a method to feed pesticide contaminated food to both an individual honey bee and a beehive colony. The procedure evaluates the pesticide effect on individual honey bees by in vivo feeding of basic larval diet and also on the natural condition of beehive colony.

The presence of pesticides in the beekeeping environment is one of the most serious problems that impacts the life of a honey bee. Pesticides can be ...

Nanopore DNA Sequencing for Metagenomic Soil Analysis

This article describes the steps for construction of a DNA library from soil, preparation and use of the nanopore flow cell, and analysis of the DNA sequences identified using computer software. Nanopore DNA sequencing is a flexible technique that allows for rapid microbial genome sequencing to identify bacterial and viral species, to characterize bacterial strains, and to detect genetic mutations that confer resistance to antibiotics. The advantages of nanopore sequencing (NS) for life sciences include its low complexity, reduced cost, and rapid real-time sequencing of purified genomic DNA, PCR amplicons, cDNA samples, or RNA. NS is an example of "strand sequencing" which involves sequencing DNA by guiding a single stranded DNA molecule through a nanopore that is inserted into a synthetic polymer membrane. The membrane has an electrical current applied across it, so as the individual bases pass through the nanopore the electrical current is disrupted to varying degrees by the four nucleotide bases. The identification of each nucleotide occurs by detecting the characteristic modulation of the electrical current by the different bases as they pass through the nanopore. The NS system consists of a handheld, USB powered portable device and a disposable flow cell that contains a nanopore array. The portable device plugs into a standard laptop computer that reads and records the DNA sequence using computer software.

Nanopore technology for sequencing biomolecules has wide applications in the life sciences, including identification of pathogens, food safety monitoring, genomic analysis, metagenomic environmental monitoring, and characterization of bacterial antibiotic resistance. In this article, the procedure for metagenomic soil DNA sequencing for species identification using the nanopore sequencing technology is demonstrated.

This article describes the steps for construction of a DNA library from soil, preparation and use of the nanopore flow cell, and analysis of the DNA ...

RGB and Spectral Root Imaging for Plant Phenotyping and Physiological Research: Experimental Setup and Imaging Protocols

Better understanding of plant root dynamics is essential to improve resource use efficiency of agricultural systems and increase the resistance of crop cultivars against environmental stresses. An experimental protocol is presented for RGB and hyperspectral imaging of root systems. The approach uses rhizoboxes where plants grow in natural soil over a longer time to observe fully developed root systems. Experimental settings are exemplified for assessing rhizobox plants under water stress and studying the role of roots. An RGB imaging setup is described for cheap and quick quantification of root development over time. Hyperspectral imaging improves root segmentation from the soil background compared to RGB color based thresholding. The particular strength of hyperspectral imaging is the acquisition of chemometric information on the root-soil system for functional understanding. This is demonstrated with high resolution water content mapping. Spectral imaging however is more complex in image acquisition, processing and analysis compared to the RGB approach. A combination of both methods can optimize a comprehensive assessment of the root system. Application examples integrating root and aboveground traits are given for the context of plant phenotyping and plant physiological research. Further improvement of root imaging can be obtained by optimizing RGB image quality with better illumination using different light sources and by extension of image analysis methods to infer on root zone properties from spectral data.

An experimental protocol is presented for assessment of soil grown plant root systems with RGB and hyperspectral imaging. Combination of RGB image time series with chemometric information from hyperspectral scans optimizes insights into plant root dynamics.

Better understanding of plant root dynamics is essential to improve resource use efficiency of agricultural systems and increase the resistance of crop ...

Techniques for Investigating the Anatomy of the Ant Visual System

This article outlines a suite of techniques in light microscopy (LM) and electron microscopy (EM) which can be used to study the internal and external eye anatomy of insects. These include traditional histological techniques optimized for work on ant eyes and adapted to work in concert with other techniques such as transmission electron microscopy (TEM) and scanning electron microscopy (SEM). These techniques, although vastly useful, can be difficult for the novice microscopist, so great emphasis has been placed in this article on troubleshooting and optimization for different specimens. We provide information on imaging techniques for the entire specimen (photo-microscopy and SEM) and discuss their advantages and disadvantages. We highlight the technique used in determining lens diameters for the entire eye and discuss new techniques for improvement. Lastly, we discuss techniques involved in preparing samples for LM and TEM, sectioning, staining, and imaging these samples. We discuss the hurdles that one might come across when preparing samples and how best to navigate around them.

This article outlines a suite of techniques in light and electron microscopy to study the internal and external eye anatomy of insects. These include several traditional techniques optimized for work on ant eyes, detailed troubleshooting, and suggestions for optimization for different specimens and regions of interest.

This article outlines a suite of techniques in light microscopy (LM) and electron microscopy (EM) which can be used to study the internal and external eye ...

Isolation of Mandibular Gland Reservoir Contents from Bornean 'Exploding Ants' (Formicidae) for Volatilome Analysis by GC-MS and MetaboliteDetector

The aim of this manuscript is to present a protocol describing the metabolomic analysis of Bornean &#39;exploding ants&#39; belonging to the Colobopsis cylindrica (COCY) group. For this purpose, the model species C. explodens is used. Ants belonging to the minor worker caste possess distinctive hypertrophied mandibular glands (MGs). In territorial combat, they use the viscous contents of their enlarged mandibular gland reservoirs (MGRs) to kill rival arthropods in characteristic suicidal &#39;explosions&#39; by voluntary rupture of the gastral integument (autothysis). We show the dissection of worker ants of this species for the isolation of the gastral portion of the wax-like MGR contents as well as listing the necessary steps required for solvent-extraction of the therein contained volatile compounds with subsequent gas chromatography-mass spectrometry (GC-MS) analysis and putative identification of metabolites contained in the extract. The dissection procedure is performed under cooled conditions and without the use of any dissection buffer solution to minimize the changes in the chemical composition of the MGR contents. After solvent-based extraction of volatile metabolites contained therein, the necessary steps for analyzing the samples via liquid-injection-GC-MS are presented. Lastly, data processing and putative metabolite identification with the use of the open-source software MetaboliteDetector is shown. With this approach, the profiling and identification of volatile metabolites in MGRs of ants belonging to the COCY group via GC-MS and the MetaboliteDetector software become possible.

Isolation of Mandibular Gland Reservoir Contents from Bornean 'Exploding Ants' Formicidae for Volatilome Analysis by GC-MS and MetaboliteDetector

Minor workers of the ant species Colobopsis explodens are dissected to isolate the wax-like content stored in their hypertrophied mandibular gland reservoirs for subsequent solvent-extraction and analysis by gas chromatography-mass spectrometry. The&#160;annotation and identification of volatile constituents using the open-source software MetaboliteDetector is also described.

The aim of this manuscript is to present a protocol describing the metabolomic analysis of Bornean &#39;exploding ants&#39; belonging to the Colobopsis ...

In Situ Hybridization Techniques for Paraffin-Embedded Adult Coral Samples

Corals are important ocean invertebrates that are critical for overall ocean health as well as human health. However, due to human impacts such as rising ocean temperatures and ocean acidification, corals are increasingly under threat. To tackle these challenges, advances in cell and molecular biology have proven to be crucial for diagnosing the health of corals. Modifying some of the techniques commonly used in human medicine could greatly improve researchers&#39; ability to treat and save corals. To address this, a protocol for in situ hybridization used primarily in human medicine and evolutionary developmental biology has been adapted for use in adult corals under stress.
The purpose of this method is to visualize the spatial expression of an RNA probe in adult coral tissue that has been embedded in paraffin and sectioned onto glass slides. This method focuses on removal of the paraffin and rehydration of the sample, pretreatment of the sample to ensure permeability of the sample, pre-hybridization incubation, hybridization of the RNA probe, and visualization of the RNA probe. This is a powerful method when using non-model organisms to discover where specific genes are expressed, and the protocol can be easily adapted for other non-model organisms. However, the method is limited in that it is primarily qualitative, because expression intensity can vary depending on the amount of time used during the visualization step and the concentration of the probe. Furthermore, patience is required, as this protocol can take up to 5 days (and in many cases, longer) depending on the probe being used. Finally, non-specific background staining is common, but this limitation can be overcome.

In Situ Hybridization Techniques for Paraffin-Embedded Adult Coral Samples

The goal of this protocol is to perform in situ hybridization on adult coral samples that have been embedded in paraffin and sectioned onto glass slides. This is a qualitative method used to visualize the spatial expression of an RNA anti-sense probe in paraffin-embedded tissues.

Corals are important ocean invertebrates that are critical for overall ocean health as well as human health. However, due to human impacts such as rising ...

Necropsy-based Wild Fish Health Assessment

Anthropogenic influences from increased nutrients and chemical contaminants, to habitat alterations and climate change, can have significant effects on fish populations. Adverse effects monitoring, utilizing biomarkers from the organismal to the molecular level, can be used to assess the cumulative effects on fishes and other organisms. Fish health has been used worldwide as an indicator of aquatic ecosystem health. The necropsy-based fish health assessment provides data on visible abnormalities and lesions, parasites, condition and organosomatic indices. These can be compared by site, season and sex, as well as temporally, to document change over time. Severity ratings can be assigned to various observations to calculate a fish health index for more quantitative assessment. A drawback of the necropsy-based assessment is that it is based on visual observations and condition factors, which are not as sensitive as tissue and subcellular biomarkers for sublethal effects. Additionally, it is rarely possible to identify causes or risk factors associated with observed abnormalities. So, for instance a raised lesion or "tumor" on the fins, lips or body surface may be a neoplasm. However, it could also be a response to a parasite, chronic inflammation or hyperplasia of normal cells in response to an irritant. Conversely, neoplasms, certain parasites, other infectious agents and many tissue changes are not visible and so may be underestimated. However, during the necropsy-based assessment, blood (plasma), tissues for histopathology (microscopic pathology), genomics and other molecular analyses, and otoliths for aging can be collected. These downstream analyses, together with geospatial analyses, habitat assessments, water quality and contaminant analyses can all be important in comprehensive ecosystem evaluations.

The health of wild fishes can be used as an indicator of aquatic ecosystem health. Necropsy-based fish health assessments provide documentation of visible lesions or abnormalities, data used to calculate condition indices as well as the opportunity to collect tissues for microscopic evaluation, gene expression and other more in-depth analyses.

Anthropogenic influences from increased nutrients and chemical contaminants, to habitat alterations and climate change, can have significant effects on ...

Measuring Hypopharyngeal Gland Acinus Size in Honey Bee (Apis mellifera) Workers

The nurse hypopharyngeal glands produce the protein fraction of the worker and royal jelly that is fed to developing larvae and queens. These paired glands that are located in the head of the bee are highly sensitive to the quantity and quality of pollen and pollen substitutes that the nurse bee consumes. The glands get smaller when nurses are fed deficient diets and are large when they are fed complete diets. Because nurse hypopharyngeal gland size is a robust indicator of nurse nutrition, it is essential that those studying honey bee nutrition know how to measure these glands. Here, we provide detailed methods for dissecting, staining, imaging, and measuring nurse bee hypopharyngeal glands. We present comparisons of unstained and stained tissue and data that were used to study the impact of pollen on gland size. This method has been used to test how diet impacts hypopharyngeal gland size but has further use for understanding the role of these glands in hive health.

Measuring Hypopharyngeal Gland Acinus Size in Honey Bee Apis mellifera Workers

Hypopharyngeal gland acinus size is a robust measure of nurse honey bee nutrition. Here, we provide a detailed protocol for dissecting, staining, imaging, and measuring nurse bee hypopharyngeal gland acini.

The nurse hypopharyngeal glands produce the protein fraction of the worker and royal jelly that is fed to developing larvae and queens. These paired ...

Empirical, Metagenomic, and Computational Techniques Illuminate the Mechanisms by which Fungicides Compromise Bee Health

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