This research was supported by funds from the Yale Climate and Energy Institute and the US National Science Foundation.

1. Rearing Grasshoppers Under Stress and Stress Free Conditions
<ol>
 <li>Use 0.5 m2 circular, closed mesocosms to prevent emigration or immigration of animal species (Figure 1). Construct mesocosms using 2.4 m lengths of 1.5 m high &frac14;" mesh aluminum fence as a scaffolding. Cover the fencing with 2.5 m lengths 1.75 m high aluminum window screening folded over the top and bottom of the fencing and stapled together along fold. Join the fencing ends to form a closed circle and then staple the overlapping window screening together to create a seal. Set the mesocosm into the soil in the field by digging a 10 cm deep by 4 cm wide trench around the base of the mesocosom, sink the mesocosm into the trench and then tamp the sod of the trench around the sunken part of the mesocosm. Staple a circular piece of window screening to the top of the mesocosm.</li>
 <li>Array mesocosms in a replicated paired experimental design in the field. Plot locations should be selected to match the plant species identity and plant relative cover. Sink cages 10 cm into the ground at the plot site.</li>
 <li>Using a sweep net, collect early (2nd) instar grasshopper nymphs and stock them into the mesocosms at natural field densities.</li>
 <li>Using a sweep net, capture individuals of a dominant sit-and-wait hunting (not web weaving) spider predator species. Glue shut the spider chelicerae (mouthparts used to subdue prey) with fast-drying cement to decoupled risk effects from actual survival selection favoring individual grasshoppers with better abilities to evade spider predation. Stock the spiders at field densities to one mesocosm of each pair. This will be the stress treatment. Mesocosms without spiders will be the stress free treatment.</li>
 <li>Allow grasshopper nymphs to develop to late (4th and 5th) instar stages. Collect all individuals from the cages and randomly assign individuals from each cage to one of three subsequent assay groups: (1) validation of physiological stress state; (2) validation of shift in body elemental stoichiometry; (3) microbial decomposition.</li>
</ol>
2. Validating Grasshopper Stress State
<ol>
 <li>Measure grasshopper standard metabolic rate (SMR), as the rate of carbon dioxide emission (<img src="/files/ftp_upload/50061/50061eq1.jpg" fo:src="/files/ftp_upload/50061/50061eq1highres.jpg"/>) in an incurrent flow-through respirometry system with an air flow rate of 200 ml/min. Remove water vapor by passing flowing air through a drying agent.</li>
 <li>Following food deprivation of 16 hr (water should be available), weigh individual grasshoppers (&plusmn;0.1 mg), and place them in transparent 50 ml (9.2 cm L x 2.0 cm D) respirometer chambers and allow them to recover from handling for at least 10 min before measurements commence.</li>
 <li>Under constant average ambient temperature (temperature &plusmn; 1% standard error variation) within the respirometer chamber, analyze respired air using an infra-red CO2 analyzer (e.g. Qubit S151- 1 ppm resolution). Measure the mean minimal steady-state <img src="/files/ftp_upload/50061/50061eq1.jpg" fo:src="/files/ftp_upload/50061/50061eq1highres.jpg"/> for 10 min.</li>
 <li>The analyzer provides fractional CO2 concentration (parts-per-million), yet SMR should be reported as a rate, so one must transformed the recordings as <img src="/files/ftp_upload/50061/50061eq2n.jpg" fo:src="/files/ftp_upload/50061/50061eq2nhighres.jpg"/> where <img src="/files/ftp_upload/50061/50061eq3n.jpg" fo:src="/files/ftp_upload/50061/50061eq3nhighres.jpg"/> <img src="/files/ftp_upload/50061/50061eq4n.jpg" fo:src="/files/ftp_upload/50061/50061eq4nhighres.jpg"/> <img src="/files/ftp_upload/50061/50061eq5n.jpg" fo:src="/files/ftp_upload/50061/50061eq5nhighres.jpg"/> <img src="/files/ftp_upload/50061/50061eq6n.jpg" fo:src="/files/ftp_upload/50061/50061eq6nhighres.jpg"/> assumed equal to 0.85 in herbivorous animals.</li>
</ol>
3. Validating Shift in Body Elemental Stoichiometry
<ol>
 <li>Evaluate Carbon:Nitrogen (C:N) content of a sample of grasshoppers collected from the field mesocosms.</li>
 <li>Reduce variation in C:N due to recent food consumption by removing grasshopper gut contents under a dissecting microscope.</li>
 <li>Freeze-dry the empty gut and body for 48 hr and then grind the individual carcass and gut to a homogeneous powder.</li>
 <li>Measure C:N contents of the powder using a CNH autoanalyzer.</li>
</ol>
4. Microbial Decomposition
<ol>
 <li>Place replicated pairs of PVC collars (15.4-cm dia., inserted ~4 cm into the soil) in the field site (Figure 3C). Remove all vegetation within them via clipping at the soil surface. These collars are used for decomposition measures. Additionally, establish a set of PVC collars across the field site to act as 13C natural-abundance controls (see below), to which neither grasshoppers nor grass-litter are added.</li>
 <li>To one collar in each pair add 2 intact, freeze-dried carcasses of grasshoppers (record the biomass added) reared with predator risk as described above using field-cages. To the other collar in each pair add 2 intact freeze-dried carcasses reared without predator risk.</li>
 <li>Cover the PVC collars with a screen to prevent grasshopper removal by scavengers from the plots and let the grasshopper carcasses decompose for 40 days.</li>
 <li>While carcasses are decomposing, label grass-litter with 13C.
 <ol>
 <li>Construct a clear plexiglass chamber (60 cm x 60 cm x 1.5 m) with an inlet and outlet valve (Figure 3B).</li>
 <li>Sink a square 60 cm x 60 cm wooden frame with a rubber seal coated with silicon grease 5 cm into the ground (Figure 3B).</li>
 <li>Slide the chamber on top of the wooden frame so that the chamber becomes sealed by the rubber (Figure 3B).</li>
 <li>Connect the chamber inlets to compressed gas cylinders containing 99 atom% 13CO2. Plants inside the chamber will be labelled with 13C, where CO2 concentrations are maintained at ambient levels (because elevating concentrations alters plant carbon partitioning). Ambient levels are maintained by only pulsing labeled CO2 for short periods of time. Also, chamber temperatures are monitored and chambers are removed if temperatures reach 5 &deg;C above ambient.</li>
 <li>One week after labeling, compare &delta;13C of the grass-litter with natural abundance values collected from a random sample of identical grass species using a Thermo DeltaPlus isotope ratio mass spectrometer (Thermo, San Jose, CA, USA).</li>
 </ol>
 </li>
 <li>After 40 days, add 10 g of air-dried 13C-labeled grass-litter to each collar that had previously been amended with grasshopper carcasses.</li>
 <li>Monitor the mineralization rate of the grass-litter in situ across 75 days by capping each collar and tracking both total soil respiration and the respiration of 13CO2. This is accomplished using a flow-through chamber technique where gas samples from each collar are monitored, in real time, for 8 min each using cavity ring-down spectroscopy (CRDS; Picarro Inc., Santa Clara, CA, USA; Model: G1101-i). CRDS enables one to simultaneously track both total and &delta;13C of soil respiration.</li>
 <li>Estimate the contribution of 13C-labeled grass-litter to total soil respiration using isotope mixing equations. The amount of grass-litter derived CO2 is calculated as follows: Cgrass-litter derived = Ctotal &times; (&delta;13Crespiration- &delta;13Cnat.abn.)/ (&delta;13Cgrass-litter- &delta;13Cnat.abn), where Ctotal is the total amount of C respired during a given measurement, &delta;13Crespiration is the &delta;13C of respired-C for collars amended with labeled grass litter, &delta;13Cnat.abn. is the mean &delta;13C of respired C in the three natural abundance collars (i.e. those that were not amended with litter), and &delta;13Cgrass-litter is the &delta;13C of the grass litter added to the collars.</li>
 <li>Monitor both soil temperature and moisture across the experiment using hand-held probes to correct for differences in soil respiration due to differences in temperature and moisture.</li>
 <li>Although intended for field use, the cavity ring-down spectroscopy instrument (Picarro Inc., Santa Clara, CA, USA; Model: G1101-i) readings are sensitive to movement. Therefore, one should erect a base measurement station central to all of the plots containing PVC collars, and connect the instrument to the collars with lengths of PVC tubing.</li>
</ol>

<ol>
	<li>Wardle, D. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ecological+linkages+between+aboveground+and+belowground+biota.">Ecological linkages between aboveground and belowground biota.</a> Science. 304, 1629-1633 (2004).</li><li>Hattenschwiler, S., Tiunov, A. V., Scheu, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biodiversity+and+litter+decomposition+in+terrestrial+ecosystems.">Biodiversity and litter decomposition in terrestrial ecosystems.</a> Annu. Rev. Ecol. Syst. 36, 191-218 (2005).</li><li>Cebrian, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+first-order+consumers+in+ecosystem+carbon+flow.">Role of first-order consumers in ecosystem carbon flow.</a> Ecol. 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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physiological+stress+as+a+fundamental+mechanism+linking+predation+to+ecosystem+functioning.">Physiological stress as a fundamental mechanism linking predation to ecosystem functioning.</a> Am. Nat. 176, 537-556 (2010).</li><li>Stoks, R. D. e. B. l. o. c. k., M, M. A., McPeek, <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Alternative+growth+and+energy+storage+responses+to+mortality+threats+in+damselflies.">Alternative growth and energy storage responses to mortality threats in damselflies.</a> Ecol. Lett. 8, 1307-1316 (2005).</li><li>Hawlena, D., Schmitz, O. 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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Periodical+cicadas+and+resource+pulses+in+North+American+forests.">Periodical cicadas and resource pulses in North American forests.</a> Science. 306, 1565 (2004).</li><li>Belovsky, G. E., Slade, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insect+herbivory+accelerates+nutrient+cycling+and+increases+plant+production.">Insect herbivory accelerates nutrient cycling and increases plant production.</a> Proc. Nat. Acad. Sci. U.S.A. 97, 14412 (2000).</li><li>Frost, C. J., Hunter, M. 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We have nothing to disclose.

The sequence of methods presented here should allow systematic measurement of the way stress in species comprising above-ground food webs can prime soil microbial communities in ways that lead to alteration of subsequent decomposition of plant litter. The methods are ideal for studying ecosystems comprised of arthropod consumers and herbaceous plants because intact food webs can be spatially circumscribed and contained within mesocosms.
Spatial variability may exist due to gradients in background soil moisture, soil temperature, plant nutrient content, etc. The study design allows one to array mescosms and PVC collars to block along spatial environmental gradients and thereby account for such environmental variation when analyzing for effects.
Although intended for field use, the cavity ring-down spectroscopy instrument (Picarro Inc., Santa Clara, CA, USA; Model: G1101-i) readings are sensitive to movement. Therefore, one should erect a base measurement station central to all of the plots containing PVC collars, and connect the instrument to the collars with lengths of PVC tubing.
Soil litter decomposition has traditionally been measured by enclosing known quantities of litter into fiberglass mesh bags, depositing the bags onto the soil surface in the field and periodically re-measuring the bags to quantify litter disappearance rate (decomposition). The limitation of this method is that one is unable to trace the fate of the decomposed matter or determine the contribution to CO2 mineralization of the soil amendment (added litter) from background soil CO2 mineralization. The tracer method using labeled CO2 presented here helps alleviate this logistical constraint.
Ecosystem ecology and biogeochemistry have operated under the working paradigm that because uneaten plant-litter comprises the majority of detritus, belowground ecosystem processes are only marginally influenced by biomass inputs from higher trophic levels in aboveground food webs, such as herbivores themselves6. However, there is growing evidence that species in higher trophic levels of ecosystems can have a profound influence on belowground processes1,4,5. The method presented here stands to enhance quantification of the contribution of higher trophic levels, either directly through biomass from carcass deposition (e.g. 12, 13) or excretion and egestion (e.g. 14, 15) or indirectly through alteration of plant community composition (e.g. 9) on ecosystem nutrient cycling. Such quantification can help reveal the mechanisms by which animals control ecosystem dynamics as part of a concerted effort to enhance and revise the current working paradigm of biotic control over ecosystem functioning.

<table border="1">
<tbody>
 <tr>
 <td>Name of the reagent or 			equipment</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Cavity ring down spectroscope</td>
 <td>Picarro Inc., Santa Clara, 			CA, USA</td>
 <td>Model # G1101-i</td>
 <td></td>
 </tr>
 <tr>
 <td>CO2 respirometer</td>
 <td>Qubit Systems, Kingston, ON, Canada</td>
 <td>Model # S151</td>
 <td></td>
 </tr>
 <tr>
 <td>13C</td>
 <td>Sigma-Aldrich</td>
 <td>372382</td>
 <td></td>
 </tr>
 <tr>
 <td>Spectrophotometer</td>
 <td>Thermo, San Jose CA, USA</td>
 <td>Model: Delta V Plus Isotope Ratio Mass Spectrophotometer</td>
 <td></td>
 </tr>
</tbody>
</table>

linking predation risk, herbivore physiological stress and microbial decomposition of plant litter

The quantity and quality of detritus entering the soil determines the rate of decomposition by microbial communities as well as recycle rates of nitrogen (N) and carbon (C) sequestration1,2. Plant litter comprises the majority of detritus3, and so it is assumed that decomposition is only marginally influenced by biomass inputs from animals such as herbivores and carnivores4,5. However, carnivores may influence microbial decomposition of plant litter via a chain of interactions in which predation risk alters the physiology of their herbivore prey that in turn alters soil microbial functioning when the herbivore carcasses are decomposed6. A physiological stress response by herbivores to the risk of predation can change the C:N elemental composition of herbivore biomass7,8,9 because stress from predation risk increases herbivore basal energy demands that in nutrient-limited systems forces herbivores to shift their consumption from N-rich resources to support growth and reproduction to C-rich carbohydrate resources to support heightened metabolism6. Herbivores have limited ability to store excess nutrients, so stressed herbivores excrete N as they increase carbohydrate-C consumption7. Ultimately, prey stressed by predation risk increase their body C:N ratio7,10, making them poorer quality resources for the soil microbial pool likely due to lower availability of labile N for microbial enzyme production6. Thus, decomposition of carcasses of stressed herbivores has a priming effect on the functioning of microbial communities that decreases subsequent ability to of microbes to decompose plant litter6,10,11.
We present the methodology to evaluate linkages between predation risk and litter decomposition by soil microbes. We describe how to: induce stress in herbivores from predation risk; measure those stress responses, and measure the consequences on microbial decomposition. We use insights from a model grassland ecosystem comprising the hunting spider predator (Pisuarina mira), a dominant grasshopper herbivore (Melanoplus femurrubrum),and a variety of grass and forb plants9.

An example plot of grasshopper standard metabolic rates in stress and stress free conditions are presented in Figure 2. Due to body mass differences among individual grasshoppers, and the fact that metabolic rate varies with body mass, plots should present metabolic rates in relation to grasshopper body mass. Parallel trends for the different treatments indicate that metabolic rate rises as a constant multiple of standard metabolic rate (i.e. there is no body mass x metabolic rate interaction) for all stressed individuals.
Grasshopper body C and N elemental contents in risk and risk free conditions are presented in Table 1. It is noteworthy that there is a very small (4%) difference in body C:N ratios between treatments. Nevertheless, these small differences can translate into large differences in grass litter decomposition by the soil microbial pool (Figure 3).
Adding grass litter to PVC collars previously amended with stressed or stress-free grasshoppers leads to different degrees of litter decomposition, as reflected in the curves describing cumulate CO2 release from the soil due to microbial respiration (Figure 3). Experiments should be monitored until cumulate curves begin to saturate.
<table border="1">
 <tbody>
 <tr>
 <td></td>
 <td>Stress</td>
 <td>Stress Free</td>
 </tr>
 <tr>
 <td>Carbon (%)</td>
 <td>48.44&plusmn;0.32</td>
 <td>44.73&plusmn;0.46</td>
 </tr>
 <tr>
 <td>Nitrogen (%)</td>
 <td>12.11&plusmn;0.08</td>
 <td>11.62&plusmn;0.12</td>
 </tr>
 <tr>
 <td>Carbon: Nitrogen</td>
 <td>4.00&plusmn;0.03</td>
 <td>3.85&plusmn;0.04</td>
 </tr>
 </tbody>
</table>
Table 1. Comparison of the chemical content of grasshopper herbivore carcasses from conditions in which they faced predation risk (stress) and in which predation risk was absent (stress free). Values are mean &plusmn; 1 standard error.
<img src="/files/ftp_upload/50061/50061fig1.jpg" alt="Figure 1"/> 
 Figure 1. Illustration of the design of the field mesocosms used in the experiment and overall scheme of the experimental evaluation of risk effects on litter decomposition.
<img src="/files/ftp_upload/50061/50061fig2.jpg" alt="Figure 2"/> 
 Figure 2. A plot of herbivore standard metabolic rate in relation to herbivore body mass. The data are divided into two classes according to experimental treatment: grasshoppers from mesocosms containing predators (predation) to induce stress, and mesocosms without predators (control) and hence no induced stress. Data are from D. Halwena and O.J. Schmitz 2010, unpublished.
<img src="/files/ftp_upload/50061/50061fig3.jpg" alt="Figure 3" fo:content-width="6in" fo:src="/files/ftp_upload/50061/50061fig3highres.jpg"/> 
 Figure 3. Curves describing cumulative CO2 release by the microbial pool while decomposing experimental grass litter inputs in PVC collars. Plotted values are mean &plusmn; 1 standard error. The graph demonstrates that soils primed with stressed grasshopper carcasses (predator) result in 19% lower (ANOVA F1,6 = 9.06, P &lt; 0.05) plant litter decomposition rates than soils primed with stress free grasshopper carcasses (control). The inset shows the PVC collar apparatus in the field. Figure reproduced from Hawlena et al.6 <a href="https://www.jove.com/files/ftp_upload/50061/50061fig3large.jpg" target="_blank"> Cick here to view larger figure</a>.

Watch this Scientific Journal Video about Linking Predation Risk, Herbivore Physiological Stress and Microbial Decomposition of Plant Litter at JoVE.com

Linking Predation Risk, Herbivore Physiological Stress and Microbial Decomposition of Plant Litter

We present methods to evaluate how predation risk can alter the chemical quality of herbivore prey by inducing dietary changes to meet demands of heightened stress, and how the decomposition of carcasses from these stressed herbivores slows subsequent plant litter decomposition by soil microbes.

The quantity and quality of detritus entering the soil  determines the rate of decomposition by microbial communities as well as  recycle rates of ...

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12.9K ビュー.  Yale University. 私たちは、捕食リスクが高まりストレスの要求を満たすために食生活の変化を誘導することによって、草食動物の餌の化学的性質を変えることができるかを評価するための方法を提示し、どのようにこれらのストレス草食動物から死体の分解は土壌微生物によって後続の植物リターの分解が遅くなります。

ビデオ: Linking Predation Risk, Herbivore Physiological Stress and Microbial Decomposition of Plant Litter

Testing the Physiological Barriers to Viral Transmission in Aphids Using Microinjection

Potato loafroll virus (PLRV), from the family Luteoviridae infects solanaceous plants. It is transmitted by aphids, primarily, the green peach aphid. When an uninfected aphid feeds on an infected plant it contracts the virus through the plant phloem. Once ingested, the virus must pass from the insect gut to the hemolymph (the insect  blood ) and then must pass through the salivary gland, in order to be transmitted back to a new plant. An aphid may take up different viruses when munching on a plant, however only a small fraction will pass through the gut and salivary gland, the two main barriers for transmission to infect more plants. In the lab, we use physalis plants to study PLRV transmission. In this host, symptoms are characterized by stunting and interveinal chlorosis (yellowing of the leaves between the veins with the veins remaining green). The video that we present demonstrates a method for performing aphid microinjection on insects that do not vector PLVR viruses and tests whether the gut is preventing viral transmission.
The video that we present demonstrates a method for performing Aphid microinjection on insects that do not vector PLVR viruses and tests whether the gut or salivary gland is preventing viral transmission.

Aphids are effective transmitters of plant viruses. Aphid microinjection of virus, the procedure we will show you today, is a technique allowing researchers to inject virus directly into the hemocoel of the aphid, bypassing the gut, one of the 2 major barriers for virus transmission in a circulative manner. The same technique is also used to inject dsRNA for RNAi.

Potato loafroll virus (PLRV), from the family Luteoviridae infects solanaceous plants. It is transmitted by aphids, primarily, the green peach aphid. When ...

Monitoring Plant Hormones During Stress Responses

Plant hormones and related signaling compounds play an important role in the regulation of plant responses to various environmental stimuli and stresses. Among the most severe stresses are insect herbivory, pathogen infection, and drought stress. For each of these stresses a specific set of hormones and/or combinations thereof are known to fine-tune the responses, thereby ensuring the plant's survival. The major hormones involved in the regulation of these responses are jasmonic acid (JA), salicylic acid (SA), and abscisic acid (ABA). To better understand the role of individual hormones as well as their potential interaction during these responses it is necessary to monitor changes in their abundance in a temporal as well as in a spatial fashion. For the easy, sensitive, and reproducible quantification of these and other signaling compounds we developed a method based on vapor phase extraction and gas chromatography/mass spectrometry (GC/MS) analysis (1, 2, 3, 4). After extracting these compounds from the plant tissue by acidic aqueous 1-propanol mixed with dichloromethane the carboxylic acid-containing compounds are methylated, volatilized under heat, and collected on a polymeric absorbent. After elution into a sample vial the analytes are separated by gas chromatography and detected by chemical ionization mass spectrometry. The use of appropriate internal standards then allows for the simple quantification by relating the peak areas of analyte and internal standard.

A simple method is provided that allows for the rapid extraction and analysis of multiple plant hormones from small tissue samples. The procedure uses vapor phase extraction as the solemn purification step. Samples are analyzed by GC/MS with chemical ionization that produces mainly (M+1)+ ions.

Plant hormones and related signaling compounds play an important role in the regulation of plant responses to various environmental stimuli and stresses. ...

OLIgo Mass Profiling (OLIMP) of Extracellular Polysaccharides

The direct contact of cells to the environment is mediated in many organisms by an extracellular matrix. One common aspect of extracellular matrices is that they contain complex sugar moieties in form of glycoproteins, proteoglycans, and/or polysaccharides. Examples include the extracellular matrix of humans and animal cells consisting mainly of fibrillar proteins and proteoglycans or the polysaccharide based cell walls of plants and fungi, and the proteoglycan/glycolipid based cell walls of bacteria. All these glycostructures play vital roles in cell-to-cell and cell-to-environment communication and signalling.
An extraordinary complex example of an extracellular matrix is present in the walls of higher plant cells. Their wall is made almost entirely of sugars, up to 75% dry weight, and consists of the most abundant biopolymers present on this planet. Therefore, research is conducted how to utilize these materials best as a carbon-neutral renewable resource to replace petrochemicals derived from fossil fuel. The main challenge for fuel conversion remains the recalcitrance of walls to enzymatic or chemical degradation due to the unique glycostructures present in this unique biocomposite.
Here, we present a method for the rapid and sensitive analysis of plant cell wall glycostructures. This method OLIgo Mass Profiling (OLIMP) is based the enzymatic release of oligosaccharides from wall materials facilitating specific glycosylhydrolases and subsequent analysis of the solubilized oligosaccharide mixtures using matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF/MS)1 (Figure 1). OLIMP requires walls of only 5000 cells for a complete analysis, can be performed on the tissue itself2, and is amenable to high-throughput analyses3. While the absolute amount of the solubilized oligosaccharides cannot be determined by OLIMP the relative abundance of the various oligosaccharide ions can be delineated from the mass spectra giving insights about the substitution-pattern of the native polysaccharide present in the wall.
OLIMP can be used to analyze a wide variety of wall polymers, limited only by the availability of specific enzymes4. For example, for the analysis of polymers present in the plant cell wall enzymes are available to analyse the hemicelluloses xyloglucan using a xyloglucanase5, 11, 12, 13, xylan using an endo-&beta;-(1-4)-xylanase 6,7, or for pectic polysaccharides using a combination of a polygalacturonase and a methylesterase 8. Furthermore, using the same principles of OLIMP glycosylhydrolase and even glycosyltransferase activities can be monitored and determined 9.

OLIgo Mass Profiling OLIMP of Extracellular Polysaccharides

A rapid way is described to gain insights into the structure of polysaccharides in an extracellular matrix. The method takes advantage of the specificity of glycosylhydrolases and the sensitivity of mass spectrometry allowing minute amounts of materials to be analyzed. This technique is adaptable to be used directly on tissue itself.

The direct contact of cells to the environment is mediated in many organisms by an extracellular matrix. One common aspect of extracellular matrices is ...

Agar-Block Microcosms for Controlled Plant Tissue Decomposition by Aerobic Fungi

The two principal methods for studying fungal biodegradation of lignocellulosic plant tissues were developed for wood preservative testing (soil-block; agar-block). It is well-accepted that soil-block microcosms yield higher decay rates, fewer moisture issues, lower variability among studies, and higher thresholds of preservative toxicity. Soil-block testing is thus the more utilized technique and has been standardized by American Society for Testing and Materials (ASTM) (method D 1413-07). The soil-block design has drawbacks, however, using locally-variable soil sources and in limiting the control of nutrients external (exogenous) to the decaying tissues. These drawbacks have emerged as a problem in applying this method to other, increasingly popular research aims. These modern aims include degrading lignocellulosics for bioenergy research, testing bioremediation of co-metabolized toxics, evaluating oxidative mechanisms, and tracking translocated elements along hyphal networks. Soil-blocks do not lend enough control in these applications. A refined agar-block approach is necessary.
Here, we use the brown rot wood-degrading fungus Serpula lacrymans to degrade wood in agar-block microcosms, using deep Petri dishes with low-calcium agar. We test the role of exogenous gypsum on decay in a time-series, to demonstrate the utility and expected variability. Blocks from a single board rip (longitudinal cut) are conditioned, weighed, autoclaved, and introduced aseptically atop plastic mesh. Fungal inoculations are at each block face, with exogenous gypsum added at interfaces. Harvests are aseptic until the final destructive harvest. These microcosms are designed to avoid block contact with agar or Petri dish walls. Condensation is minimized during plate pours and during incubation. Finally, inoculum/gypsum/wood spacing is minimized but without allowing contact. These less technical aspects of agar-block design are also the most common causes of failure and the key source of variability among studies. Video publication is therefore useful in this case, and we demonstrate low-variability, high-quality results.

This video demonstrates a controlled environment approach to study degradation of lignocellulosic plant tissues by aerobic fungi. The ability to control nutrient sources and moisture is a key advantage of agar-block microcosms, but the approach often yields mixed success. We address critical pitfalls to yield reproducible, low-variability results.

The two principal methods for studying fungal biodegradation of lignocellulosic plant tissues were developed for wood preservative testing (soil-block; ...

Herbivore-induced Blueberry Volatiles and Intra-plant Signaling

Herbivore-induced plant volatiles (HIPVs) are commonly emitted from plants after herbivore attack1,2. These HIPVs are mainly regulated by the defensive plant hormone jasmonic acid (JA) and its volatile derivative methyl jasmonate (MeJA)3,4,5. Over the past 3 decades researchers have documented that HIPVs can repel or attract herbivores, attract the natural enemies of herbivores, and in some cases they can induce or prime plant defenses prior to herbivore attack. In a recent paper6, I reported that feeding by gypsy moth caterpillars, exogenous MeJA application, and mechanical damage induce the emissions of volatiles from blueberry plants, albeit differently. In addition, blueberry branches respond to HIPVs emitted from neighboring branches of the same plant by increasing the levels of JA and resistance to herbivores (i.e., direct plant defenses), and by priming volatile emissions (i.e., indirect plant defenses). Similar findings have been reported recently for sagebrush7, poplar8, and lima beans9..
Here, I describe a push-pull method for collecting blueberry volatiles induced by herbivore (gypsy moth) feeding, exogenous MeJA application, and mechanical damage. The volatile collection unit consists of a 4 L volatile collection chamber, a 2-piece guillotine, an air delivery system that purifies incoming air, and a vacuum system connected to a trap filled with Super-Q adsorbent to collect volatiles5,6,10. Volatiles collected in Super-Q traps are eluted with dichloromethane and then separated and quantified using Gas Chromatography (GC). This volatile collection method was used n my study6 to investigate the volatile response of undamaged branches to exposure to volatiles from herbivore-damaged branches within blueberry plants. These methods are described here. Briefly, undamaged blueberry branches are exposed to HIPVs from neighboring branches within the same plant. Using the same techniques described above, volatiles emitted from branches after exposure to HIPVs are collected and analyzed.

A push-pull method for collecting plant volatiles is described. The method allows for a comparison of volatiles induced by herbivore feeding, exogenous methyl jasmonate, and mechanical damage. This technique is also used to investigate the volatile response of undamaged branches to exposure to volatiles from herbivore-damaged branches within blueberry plants.

Herbivore-induced plant volatiles (HIPVs) are commonly emitted from plants after herbivore attack1,2.  These HIPVs are mainly regulated by the defensive ...

Physiological Recordings and RNA Sequencing of the Gustatory Appendages of the Yellow-fever Mosquito Aedes aegypti

Electrophysiological recording of action potentials from sensory neurons of mosquitoes provides investigators a glimpse into the chemical perception of these disease vectors. We have recently identified a bitter sensing neuron in the labellum of female Aedes aegypti that responds to DEET and other repellents, as well as bitter quinine, through direct electrophysiological investigation. These gustatory receptor neuron responses prompted our sequencing of total mRNA from both male and female labella and tarsi samples to elucidate the putative chemoreception genes expressed in these contact chemoreception tissues. Samples of tarsi were divided into pro-, meso- and metathoracic subtypes for both sexes. We then validated our dataset by conducting qRT-PCR on the same tissue samples and used statistical methods to compare results between the two methods. Studies addressing molecular function may now target specific genes to determine those involved in repellent perception by mosquitoes. These receptor pathways may be used to screen novel repellents towards disruption of host-seeking behavior to curb the spread of harmful viruses.

Physiological Recordings and RNA Sequencing of the Gustatory Appendages of the Yellow-fever Mosquito Aedes aegypti

Using two methods to estimate gene expression in the major gustatory appendages of Aedes aegypti, we have identified the set of genes putatively underlying the neuronal responses to bitter and repulsive compounds, as determined by electrophysiological examination.

Electrophysiological recording of action potentials from sensory neurons of mosquitoes provides investigators a glimpse into the chemical perception of ...

Visualizing Stromule Frequency with Fluorescence Microscopy

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but whose functions remain mysterious. Alongside growing attention on the role of chloroplasts in coordinating plant responses to stress, interest in stromules and their relationship to chloroplast signaling dynamics has increased in recent years, aided by advances in fluorescence microscopy and protein fluorophores that allow for rapid, accurate visualization of stromule dynamics. Here, we provide detailed protocols to assay stromule frequency in the epidermal chloroplasts of Nicotiana benthamiana, an excellent model system for investigating chloroplast stromule biology. We also provide methods for visualizing chloroplast stromules in vitro by extracting chloroplasts from leaves. Finally, we outline sampling strategies and statistical approaches to analyze differences in stromule frequencies in response to stimuli, such as environmental stress, chemical treatments, or gene silencing. Researchers can use these protocols as a starting point to develop new methods for innovative experiments to explore how and why chloroplasts make stromules.

Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but ...

Measurements of Physiological Stress Responses in C. Elegans

Organisms are often exposed to fluctuating environments and changes in intracellular homeostasis, which can have detrimental effects on their proteome and physiology. Thus, organisms have evolved targeted and specific stress responses dedicated to repair damage and maintain homeostasis. These mechanisms include the unfolded protein response of the endoplasmic reticulum (UPRER), the unfolded protein response of the mitochondria (UPRMT), the heat shock response (HSR), and the oxidative stress response (OxSR). The protocols presented here describe methods to detect and characterize the activation of these pathways and their physiological consequences in the nematode, C. elegans. First, the use of pathway-specific fluorescent transcriptional reporters is described for rapid cellular characterization, drug screening, or large-scale genetic screening (e.g., RNAi or mutant libraries). In addition, complementary, robust physiological assays are described, which can be used to directly assess sensitivity of animals to specific stressors, serving as functional validation of the transcriptional reporters. Together, these methods allow for rapid characterization of the cellular and physiological effects of internal and external proteotoxic perturbations.

Measurements of Physiological Stress Responses in C. Elegans

Here, we characterize cellular proteotoxic stress responses in the nematode C. elegans by measuring the activation of fluorescent transcriptional reporters and assaying sensitivity to physiological stress.

Organisms are often exposed to fluctuating environments and changes in intracellular homeostasis, which can have detrimental effects on their proteome and ...

Combining Clearing and Fluorescence Microscopy for Visualising Changes in Gene Expression and Physiological Responses to Plasmodiophora brassicae

Infection of Brassica crops by the soilborne protist Plasmodiophora brassicae leads to gall formation on the underground organs. The formation of galls requires cellular reprogramming and changes in the metabolism of the infected plant. This is necessary to establish a pathogen-oriented physiological sink toward which the host nutrients are redirected. For a complete understanding of this particular plant-pathogen interaction and the mechanisms by which host growth and development are subverted and repatterned, it is essential to track and observe the internal changes accompanying gall formation with cellular resolution. Methods combining fluorescent stains and fluorescent proteins are often employed to study anatomical and physiological responses in plants. Unfortunately, the large size of galls and their low transparency act as major hurdles in performing whole-mount observations under the microscope. Moreover, low transparency limits the employment of fluorescence microscopy to study clubroot disease progression and gall formation. This article presents an optimized method for fixing and clearing galls to facilitate epifluorescence and confocal microscopy for inspecting P. brassicae-infected galls. A tissue-clearing protocol for rapid optical clearing was used followed by vibratome sectioning to detect anatomical changes and localize gene expression with promoter fusions and reporter lines tagged with fluorescent proteins. This method will prove useful for studying cellular and physiological responses in other pathogen-triggered structures in plants, such as nematode-induced syncytia and root knots, as well as leaf galls and deformations caused by insects.

Combining Clearing and Fluorescence Microscopy for Visualising Changes in Gene Expression and Physiological Responses to Plasmodiophora brassicae

The present protocol describes an optimized method for the histological observation of galls induced by Plasmodiophora brassicae. Vibratome sections of hypocotyls are cleared before fluorescence imaging to study the involvement of transcription factors and phytohormones during disease progression. This protocol overcomes resin embedding limitations, enabling in planta visualization of fluorescent proteins.

Infection of Brassica crops by the soilborne protist Plasmodiophora brassicae leads to gall formation on the underground organs. The formation of galls ...

Scalable, Flexible, and Cost-Effective Seedling Grafting

Early-stage seedling grafting has become a popular tool in molecular genetics to study root-shoot relationships within plants. Grafting early-stage seedlings of the small model plant, Arabidopsis thaliana, is technically challenging and time consuming due to the size and fragility of its seedlings. A growing collection of published methods describe this technique with varying success rates, difficulty, and associated costs. This paper describes a simple procedure to make an in-house reusable grafting device using silicone elastomer mix, and how to use this device for seedling grafting. At the time of this publication, each reusable grafting device costs only $0.47 in consumable materials to produce. Using this method, beginners can have their first successfully grafted seedlings in less than 3 weeks from start to finish. This highly accessible procedure will allow plant molecular genetics labs to establish seedling grafting as a normal part of their experimental process. Due to the full control users have in the creation and design of these grafting devices, this technique could be easily adjusted for use in larger plants, such as tomato or tobacco, if desired.

This protocol describes a robust seedling grafting method that requires no prior experience or training and can be executed at a very low cost using materials easily accessible in most molecular biology labs.

Early-stage seedling grafting has become a popular tool in molecular genetics to study root-shoot relationships within plants. Grafting early-stage ...