I thank the following individuals for their assistance in the development of this protocol: Chuck Kristensen, Greta Binford, Alex K. Lancaster, Konrad Zinsmaier, and Mays Imad. Mass spectrometry and proteomics data were acquired by the Arizona Proteomics Consortium supported by NIEHS grant ES06694 to the SWEHSC, NIH/NCI grant CA023074 to the AZCC and by the BIO5 Institute of the University of Arizona. Funding for this work was provided from the National Institutes of Health (National Institute of General Medicine) from grants 1F32GM83661-01 and 1R15GM097714-01 to Jessica E. Garb.

1. Preparation of Electro-stimulator Apparatus
<ol>
	<li>Connect electro-stimulator power cord to outlet and attach external foot pedal to the external signal input. 
	Note: Electrical wires must be properly insulated and exposed metal delivering current should not be touched. Wear latex or nitrile gloves for protection. Electro-stimulator power switch should be in OFF position when attaching power cord and attaching wires.</li>
	<li>Connect positive electrode to red output terminal on electro-stimulator (when polarity switch is in normal mode) and connect the negative electrode to the black output terminal. Note: Each electrode wire should terminate in an alligator clip.</li>
	<li>Set stimulator to the following recommended initial settings: voltage=7 V, Frequency=1 pulses per second, delay=0 milliseconds, duration=200 milliseconds, twin pulses switch=regular, and mode switch=off.</li>
	<li>Test output of electrodes by attaching alligator clips to voltmeter positive and negative test leads. Turn on stimulator power switch and deliver current with foot pedal.</li>
</ol>
2. Preparation of Immobilizing Forceps and Other Materials
<ol>
	<li>Dip one prong of featherweight forceps into liquid plastic coating and wait four hours to let coating dry. Tightly wrap 15 mm of the tip of the opposing prong with a single layer of cotton sewing thread, and secure thread by tying the end in a knot. 
	Note: The thread is used to absorb an applied saline solution that promotes electrical conductivity, while the plastic coating provides insulation to retard current.</li>
	<li>Cut tip of blunt hypodermic needle with sharp scissors and smooth the opening with sand paper. Attach the needle to a vacuum waste trap (e.g., vacuum filter flask) via tubing. 
	Note: &#160;The needle will suction away water and regurgitate, and will serve to complete the circuit created by the stimulator electrodes. The needle opening must be small enough to not harm the spider, yet large enough to vacuum solution without clogging (e.g., 25 G x 1 &#189; in gauge, see Materials List).</li>
	<li>Position featherweight forceps horizontally in a tightly closed position using a vinyl-covered two-pronged clamp secured to a magnetic base or fixed stand apparatus under a dissecting microscope&#8217;s field of view, with plastic coated prong of forceps on top and thread coated prong on bottom. 
	Note: The two-pronged clamp is maintained in a stationary position to the magnetic base by a clamp holder (see Materials List). A rubber band may be tightly secured around prongs of the featherweight forceps, next to the clamp holder, to add additional pressure to keep the featherweight forceps in a closed position around the spider once introduced.</li>
	<li>Attach positive electrode alligator clip to forceps&#8217; bottom prong upstream of thread and attach negative alligator clip to blunt metal vacuum needle.&#160;</li>
	<li>Test circuit current with voltmeter by placing positive lead on forceps prong between thread and alligator clip and placing negative lead on blunt vacuum needle. Press foot pedal to ensure sufficient current is detected and remove leads if sufficient current is detected.</li>
	<li>Prepare several microcapillary tubes for venom collection. Hold a single microcapillary tube at one end with a gloved hand and at the other end with sterile metal forceps, placing the forceps-held end horizontally over a Bunsen burner flame. Pull on the microcapillary with the metal forceps away from the flame to create an elongated tip, while holding the other end in a stationary position. 
	Note: wear protective eyewear as microcapillaries can easily fracture.</li>
	<li>Rest prepared microcapillaries on a mounting putty strip in a petri dish. Examine microcapillary ends under a dissecting microscope and create a small, beveled opening at the pulled end with sterilized metal forceps.</li>
	<li>Label variable number of sterile 0.5 ml tubes. Add sterile, deionized water to sterile tubes (e.g., 5 &#956;l of autoclaved deionized water). Close tubes and place tubes on ice.</li>
	<li>Prepare saline solution in a beaker and fill a second beaker with autoclaved water.</li>
</ol>
3. Immobilization of Spider in Forceps
<ol>
	<li>Turn on CO2 tank and transfer spider from closed plastic collecting vial (see Materials List) into CO2 chamber using long metal forceps. 
	Note: black widow spiders have hazardous venom; always wear gloves during this protocol.</li>
	<li>Keep spider in chamber for 5-10 min or until anesthetized and no longer moving when gently prodded with long forceps. Use transfer pipette to wet thread on forceps with saline solution. 
	Note: For guidelines on CO2 anesthesia parameters for black widows, refer to Spagna and Moore19.</li>
	<li>Retrieve anesthetized spider from CO2 chamber by picking it up by its anterior legs using blunt dissecting forceps and gloved hands. Move anesthetized spider to featherweight forceps apparatus. Separate prongs of closed featherweight forceps and place the spider&#8217;s cephalothorax in between the prongs and allow the prongs to close shut to ensure spider is held tightly in place but is not being crushed. Ensure that the spider is placed carapace dorsum side downwards (see Foelix20 for guide to spider anatomy), resting on thread coated prong, and the carapace&#8217;s ventral side facing the plastic coating, while the positive electrode remains attached to bottom prong. Work quickly when handling spider.</li>
	<li>Adjust dissecting microscope magnification, focus, and light source until the spider&#8217;s chelicerae and fangs are clearly visible.</li>
</ol>
4. Venom Collection
<ol>
	<li>Turn on vacuum and spray spider chelicerae with water solution using a second blunt tipped syringe needle. Suction away water with vacuum needle to remove impurities.</li>
	<li>Touch the vacuum needle with attached negative electrode to spider rostrum and deliver pulse with foot pedal one or more times. Vacuum away regurgitate if necessary. 
	Note: spider rostrum is inside mouth posterior to chelicerae on dorsal surface between chelicerae and endites (see Foelix20 for detailed spider anatomy). Note: With each pulse, the spider&#8217;s legs contract, and venom will be visible as clear droplets emerging from fangs.</li>
	<li>Collect venom droplets with microcapillary tip and transfer capillary tip into 0.5 ml tube with water or buffer solution (solution will be sucked into capillary). 
	Note: avoid puncturing spider with capillary which may lead to hemolymph contamination and death. Also avoid contamination of capillary with silk and glue from spinnerets.</li>
	<li>Attach transfer syringe with tube adapter to microcapillary (at end opposite to collection tip) and dispense venom solution back into tube. Put tube on ice. 
	Note: Dispose of waste capillaries in a nearby sharps container.</li>
	<li>Place open plastic collecting vial, along with its cap, close to the spider. Carefully grasp the spider legs with blunt dissecting forceps held in one hand and carefully open featherweight forceps prongs with other hand, sliding spider into collecting vial with blunt forceps and quickly close cap. Continue with next spider and store venom-containing tubes in -80 &#176;C freezer when finished. 
	Note: continue to exercise caution when moving the spider from the forceps back into the vial. Make sure the collecting vial and cap is nearby to the spider for its rapid transfer and wear gloves.</li>
</ol>
5. Venom Gland Dissections
<ol>
	<li>Two-to-three days after venom collection, anesthetize spider in CO2 chamber (see step 3.1). Fill liquid nitrogen carrier and place next to dissecting microscope. 
	Note: when working with liquid nitrogen, use eye protection and cryogenic gloves.</li>
	<li>Clean dissection area and dissecting forceps with solution that eliminates RNase and DNA contamination. Fill a small petri dish placed under dissecting microscope with dissecting buffer (e.g., 1x saline-sodium citrate (SSC) buffer).&#160;</li>
	<li>Transfer spider from CO2 chamber to petri dish and use forceps to quickly separate the cephalothorax from the abdomen.</li>
	<li>Hold the cephalothorax under the dissecting microscope with forceps and position the chelicerae into field of view. Use the sharp ends of a second pair of forceps to cut cuticle joining the carapace to the lateral aspects of the chelicerae. Laterally grasp the chelicerae using the second pair of forceps, and gently tug back and forth until venom glands are pulled out.&#160;</li>
	<li>Separate glands from the chelicerae (or leave attached to chelicerae if desired) and transfer to a cyrosafe 0.5 ml tube. Place closed tube in liquid nitrogen. 
	Note: each dissection should take no more than 15 min.</li>
	<li>Repeat dissection with additional spiders until finished. Transfer tubes from liquid nitrogen into -80 &#176;C freezer.</li>
</ol>

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M., Bulaj, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuroprotective+and+cardioprotective+conopeptides:+An+emerging+class+of+drug+leads.">Neuroprotective and cardioprotective conopeptides: An emerging class of drug leads.</a> Curr. Opin. Drug Disc. 12 (2), 231-239 (2009).</li><li>Casewell, N. R., W&#252;ster, W., Vonk, F. J., Harrison, R. A., Fry, B. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Complex+cocktails:+the+evolutionary+novelty+of+venoms.">Complex cocktails: the evolutionary novelty of venoms.</a> Trends Ecol. Evol. 28 (4), 219-229 (2013).</li><li>Vetter, R. S., Isbister, G. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Medical+aspects+of+spider+bites.">Medical aspects of spider bites.</a> Annu. Rev. Entomol. 53, 409-429 (2008).</li><li>Adams, M. E., Myers, R. A., Imperial, J. S., Olivera, B. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Toxityping+rat+brain+calcium+channels+with+omega-toxins+from+spider+and+cone+snail+venoms.">Toxityping rat brain calcium channels with omega-toxins from spider and cone snail venoms.</a> Biochemistry. 32 (47), 12566-12570 (1993).</li><li>Silva, J. -. P., 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=Latrophilin+1+and+its+endogenous+ligand+Lasso/Teneurin-2+form+a+high-affinity+transsynaptic+receptor+pair+with+signaling+capabilities.">Latrophilin 1 and its endogenous ligand Lasso/Teneurin-2 form a high-affinity transsynaptic receptor pair with signaling capabilities.</a> PNAS. 108 (29), 12113-12118 (2011).</li><li>Williams, J. A., Day, M., Heavner, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ziconotide:+an+update+and+review.">Ziconotide: an update and review.</a> Expert Opin. 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The author has nothing to disclose and no competing financial interests.

Venoms represent an important source of physiologically reactive proteins, peptides and other molecules with applications for drug discovery, as well as for fundamental aspects of cellular and ecological research1&#8211;3. However, the collection of venom, particularly from dangerous or small animals, is a challenging task. This protocol demonstrates how venom and venom glands can be collected from black widow spiders, and confirms the success of this approach via a combination of MuDPIT analysis of the venom and a protein database derived from cDNAs cloned from venom glands21. While this protocol works well for black widows and medium sized spiders, other venom collection techniques have been employed for larger mygalomorph (tarantula-like) spiders, such as direct aspiration of venom from fangs into glass pipettes e.g., 24. This latter approach, however, will not work well for smaller-sized spiders that are not aggressive.&#160;

One especially critical aspect of the venom collection protocol described here is the initial phases of preparation and optimization of the collection process so that it becomes more routine, consistent and quicker. The protocol is initially challenging to master, but with repeated trials, it becomes easier and faster. Caution is also urged at all critical phases involving the handling of hazardous spiders, the use of electrical current, fine-point glass micro capillaries, and syringe needles. It is important to wear appropriate personal protective equipment such as nitrile gloves, a lab coat, long pants and closed shoes, as well as eyewear when shaping micro capillaries.&#160;

Another challenging aspect to venom collection is the small amount of venom produced by any one spider, particularly Latrodectus species, from which the amounts collected may be limited to 1-2 microliters per individual at best. Obtaining sufficient venom for downstream applications, such as protein gels or functional assays may require the combination of venom from multiple individuals into one tube. In such cases, venom should only be combined from individuals of the same sex, ontogenetic stage, and population given the recognition of intersexual, developmental and geographic variability in some venoms25,26. Spiders may also exhibit considerable variation in the amount of venom produced among individuals, where lesser amounts may reflect the recent depletion of the gland. Thus it may be advisable to collect venom several days after their last feeding. If little venom is released, excessive current should not applied to the spider, which may cause the cuticle to rupture, leading to contamination of the venom with hemolymph or death.&#160;

Contamination of venom samples with spider silk or human sources should also be avoided through the use of sterile or clean equipment. Despite these challenges, collection of pure venom, leaving the spider alive, is preferable to methods that obtain venom from gland homogenates (which do not separate venom components from other cellular proteins) and kill the spider. It is also critical to ensure that samples are quickly frozen to prevent protein degradation.

The extraction of venom promotes subsequent venom production, thereby stimulating venom gene expression in the venom gland. Thus, because this protocol allows for spiders to survive venom depletion, their glands may be dissected several days later (killing the spider) at a point where venom gene expression is expected to be sufficient for genetic studies, such as transcript cloning10,11. Several important precautions must also be taken in venom gland dissections. Emphasis should be placed on using lab equipment and reagents that are free of RNases that degrade RNA. Thus it is recommended to wipe forceps and other non-disposable equipment and surfaces with solutions that eliminate RNase and DNA contamination. The dissections should be performed as quickly as possible and directly frozen to further ensure RNA integrity of the tissue. Finally, dissections should only be performed on anesthetized spiders, after their cephalothorax and abdomen are quickly separated.

In conclusion, this article provides a verified protocol to obtain spider venom and venom glands. Venom and venom glands allow for the isolation and characterization of their protein and peptide components using proteomic and transcriptomic approaches. In addition, venom samples may represent the starting point of functional assays, which determine the biomedical and pharmacological potential of their constituent molecules. Nearly all spiders produce venom, and the wide diversity of venom components synthesized by individual species suggests a vast diversity of venom molecules are yet to be discovered13. Accordingly, this protocol provides tools to investigate the rich source of biologically active molecules present in spider venoms.

Venoms are animal secretions predominantly injected into another animal for the purposes of predation or defense, and also have important biological applications with biomedical relevance1&#8211;3. Only certain animals synthesize venom, but its production is taxonomically widespread across invertebrates (e.g., cnidarians, cone snails, scorpions, and spiders) and vertebrates (e.g., snakes, some fish and mammals), because it has independently evolved multiple times1,4. Biochemical characterization of venoms typically show they are composed of a wide variety of proteins and peptides, which largely act on the circulatory and nervous systems of injected animals to rapidly deliver constituent toxins1. A small fraction of venomous animals may pose a threat to humans, particularly species with synanthropic distributions5. The study of venom composition and functional activities has played an important role in the functional characterization of vertebrate cellular components (particularly neuronal ion channels)6 and in elucidating fundamental cellular processes (e.g. neurosecretion)7. Moreover, select venom peptides have useful properties in biomedical contexts, including their uses as treatments for cancer and pain8,9, and are mined for candidate drug leads and antivenom development. Venoms also play prominent roles in ecological and evolutionary research1,4,10,11, thus the collection of these hazardous secretions has many utilities.

There are &#62;40,000 described species of spider (Order Araneae), and all but one of the more than 100 spider families possess paired venom glands that terminate in fangs12. The high species diversity of spiders suggests that they represent the largest clade of venomous organisms. However, biochemical characterization of spider venoms has largely concentrated on a small number of species most often associated with human envenomation. Recent proteomic and transcriptomic studies of spider venoms indicate they typically contain many unique proteins and peptides2,13,14. Advances in high-throughput cDNA sequencing and mass spectrometry peptide fingerprinting has greatly facilitated the discovery of these venom proteins15. Nevertheless, such work begins with the collection of sufficient venom and/or venom glands from the spiders, and detailed documentation for such techniques are few16.

This paper presents a protocol for the collection of venom and venom glands from black widow spiders, which can be applied to similarly sized spiders. The collection of venom separately from the glands enables identification of proteins that are secreted into the venom as opposed to proteins performing other cellular functions. Black widows and other Latrodectus species are widely recognized as being among the most hazardous spiders due to their highly neurotoxic venom, which causes severe pain in humans, which is sometimes accompanied by profuse sweating, muscle contractions, hypertension, difficulty breathing and patchy paralysis5. The venom collection protocol presented here uses electro-stimulation to deliver electrical current to anesthetized spiders to elicit muscular contractions and release of venom. Venom droplets are quickly collected with micro-capillaries and dispensed into tubes for freezer storage. Because the protocol involves hazardous procedures, it should only be performed by well-trained individuals and caution is urged at key steps. The collected venom has numerous uses, such as characterization and isolation of constituent molecules2, for physiological experiments or functional assays18, and to stimulate venom gene expression11. The protocol concludes with a description of venom gland dissection and preservation useful for the cloning of venom-specific genes, the expression of which has been shown to occur 2-3 days following venom depletion in different spiders10,11.

<table><tbody><tr><td>Nerve and muscle stimulator (electro-stimulator)&amp;nbsp;</td><td>Grass Technologies</td><td>SD9</td><td>http://www.grasstechnologies.com/products/stimulators/stimsd9.html</td></tr><tr><td>Voltmeter</td><td>RadioShack</td><td>22-223</td><td>any generic voltmeter/multimeter can be substituted</td></tr><tr><td>Pointed Featherweight Forceps</td><td>Bioquip</td><td>4748</td><td></td></tr><tr><td>Plasti Dip</td><td>Performix</td><td></td><td>Available at Ace Hardware</td></tr><tr><td>25 G x 1&amp;#189; in Precision Glide hypodermic syringe needle</td><td>BD Medical</td><td>305127</td><td></td></tr><tr><td>Vacuum filter flask 1 L</td><td>Nalgene</td><td>DS4101-1000</td><td>smaller flask sizes may also work</td></tr><tr><td>Buchner Two Piece funnel, 90 mm</td><td>Nalgene</td><td>4280-0900</td><td></td></tr><tr><td>5 &amp;#956;l Capillary Bores (Micro capillaries)</td><td>VWR</td><td>53508-375</td><td></td></tr><tr><td>Mounting putty strip&amp;nbsp;</td><td>Loctite</td><td></td><td>Available at Ace Hardware</td></tr><tr><td>Fisherbrand General-Purpose Extra-Long Forceps Length: 11-13/16 in.</td><td>Fisher</td><td>10-316C</td><td></td></tr><tr><td>SSC Buffer, 20x (pH 7.0), Molecular Grade</td><td>Promega</td><td>V4261</td><td>can be made from stock chemicals (150 mM NaCl, 15 mM sodium citrate)</td></tr><tr><td>Eppendorf safe-lock tubes 0.5 ml tubes (cryogenic safe)</td><td>Eppendorf</td><td>22363611</td><td></td></tr><tr><td>Nalgene Dewar, 1 L, HDPE, Liquid nitrogen benchtop flask</td><td>Thermo Scientific</td><td>4150-1000</td><td></td></tr><tr><td>Rnase Away</td><td>VWR</td><td>53225-514</td><td></td></tr><tr><td>Ultra Fine Tweezers (dissecting forceps)</td><td>EMS</td><td>78310-0</td><td>similar high-quality fine point forceps can be substituted</td></tr><tr><td>Foot switch/pedal</td><td>Linemaster Switch Corp.</td><td>491-S</td><td></td></tr><tr><td>Two-pronged extension clamp, with vinyl covered sleeves, 8.5 inches</td><td>VWR</td><td>21570-007</td><td>similar models could be substituted</td></tr><tr><td>Clamp Holder</td><td>VWR</td><td>89084-746</td><td>similar models could be substituted, must accommodate diameter of extension clamp rod</td></tr><tr><td>Magnetic base (holds extension clamp via clamp holder)</td><td>VWR</td><td>300042-270</td><td>similar apparatus able to securely hold extension clamp in fixed position may be substituted</td></tr><tr><td>Plastic Collecting Vials (large/40 dram)</td><td>Bioquip</td><td>8940</td><td></td></tr><tr><td>Cotton sewing thread</td><td>Joann fabric and craft store</td><td>7245855</td><td>similar product could be substituted</td></tr></tbody></table>

extraction of venom and venom gland microdissections from spiders for proteomic and transcriptomic analyses

Venoms are chemically complex secretions typically comprising numerous proteins and peptides with varied physiological activities. Functional characterization of venom proteins has important biomedical applications, including the identification of drug leads or probes for cellular receptors. Spiders are the most species rich clade of venomous organisms, but the venoms of only a few species are well-understood, in part due to the difficulty associated with collecting minute quantities of venom from small animals. This paper presents a protocol for the collection of venom from spiders using electrical stimulation, demonstrating the procedure on the Western black widow (Latrodectus hesperus). The collected venom is useful for varied downstream analyses including direct protein identification via mass spectrometry, functional assays, and stimulation of venom gene expression for transcriptomic studies. This technique has the advantage over protocols that isolate venom from whole gland homogenates, which do not separate genuine venom components from cellular proteins that are not secreted as part of the venom. Representative results demonstrate the detection of known venom peptides from the collected sample using mass spectrometry. The venom collection procedure is followed by a protocol for dissecting spider venom glands, with results demonstrating that this leads to the characterization of venom-expressed proteins and peptides at the sequence level.

The collection of venom and venom gland dissections are frequently performed to characterize venom proteins and peptides at the sequence level10,11,15. Collected venom may also be used in physiological assays to determine their functional activities18. The collection of venom will stimulate venom gland gene expression, thereby facilitating the cloning of specific toxin transcripts via RT-PCR10,11. Identification of venom proteins can also be accomplished in a high-throughput manner by integrating standard mass spectrometry techniques with sequence databases generated from venom gland cDNA libraries15,21. An example of such work, starting from venom and glands collected using the protocol detailed above, demonstrating its effectiveness, is illustrated in Figure 1.
Venom collected from L. hesperus adult females was digested with trypsin and subjected to an in solution MuDPIT analysis, which links HPLC (high performance liquid chromatography) to tandem mass spectrometry (MS/MS). Here the MuDPIT analysis was performed by the University of Arizona Proteomics Consortium. Masses of detected peptides and their dissociated fragments (daughter ions, inferred from spectra) were compared to peptide sequences theoretically predicted from translations of cDNA sequences obtained from a gene expression library constructed from L. hesperus venom glands (glands were acquired by the dissection protocol described above)21. In this experiment, 36 proteins were detected from all collected spectra, at the 99.9% probability threshold, and contained at least one detected peptide. One of the detected proteins is supported by 43 total spectra (exemplar spectra shown in Figure 1A), corresponding to three exclusive peptides, covering 35% of the protein sequence (Figure 1B). The detected protein (translated from a collected cDNA) has a top BLASTp hit to latrodectin from L. tredecimguttatus (GenBank Accession P49125.1), with an e-score of 2e-46, and shares 80% identity at the amino acid sequence level with the protein detected in this experiment. Latrodectin, which is also known as alpha-latrotoxin low molecular weight protein, is a recognized venom component of black widow spiders22,23, verifying the effectiveness of the presented protocol. Some proteins identified from the venom correspond to sequences for which no BLAST hit is found. It is unclear whether such a result is due to the limited genomic resources available for spiders as well as the limited functional information for their proteins, or because the unknown protein represents a venom contaminant. Nevertheless, gene or protein expression analyses examining the abundance of such a novel protein in venom glands relative to other tissues should reveal whether it is a genuine venom component.
<img alt="Figure 1" fo:content-width="6in" src="/files/ftp_upload/51618/51618fig1highres.jpg" width="600" /> 
Figure 1: Latrodectin peptide identified from black widow venom using mass spectrometry. (A) Representative spectra (one of 43) detected via MS/MS portion of MuDPIT analysis of L. hesperus venom which was assigned to predicted translation of a collected cDNA sequence shown in part B. Spectra shows mass to charge ratio (m/z) of detected daughter ions on the horizontal axis produced by fragmenting parent peptide (CGEEDFGEEIVK), with letters at top indicating the peptide sequence based on daughter ion masses. (B) Protein sequence to which spectra in part A was assigned, showing in red, bold and underlined text the corresponding sequence from the spectra shown in part A, additional red text (not bold) represents another peptide in this protein detected with other collected spectra. The protein sequence is translated from a cDNA sequence obtained from dissected venom glands.&#160;

Watch this Scientific Journal Video about Extraction of Venom and Venom Gland Microdissections from Spiders for Proteomic and Transcriptomic Analyses at JoVE.com

Extraction of Venom and Venom Gland Microdissections from Spiders for Proteomic and Transcriptomic Analyses

This article provides a protocol for the extraction of venom from spiders using electrical stimulation in order to 1) conduct proteomic characterization, 2) stimulate venom gland gene expression, and 3) perform functional studies of venoms. This is followed by a description of venom gland microdissections for gene expression studies.

Venoms are chemically complex secretions typically comprising numerous proteins and peptides with varied physiological activities. Functional ...

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33.3K Views.  University of Massachusetts Lowell. This article provides a protocol for the extraction of venom from spiders using electrical stimulation in order to 1) conduct proteomic characterization, 2) stimulate venom gland gene expression, and 3) perform functional studies of venoms. This is followed by a description of venom gland microdissections for gene expression studies.

Video: Extraction of Venom and Venom Gland Microdissections from Spiders for Proteomic and Transcriptomic Analyses

Electrophoretic Separation of Proteins

Electrophoresis is used to separate complex mixtures of proteins (e.g., from cells, subcellular fractions, column fractions, or immunoprecipitates), to investigate subunit compositions, and to verify homogeneity of protein samples. It can also serve to purify proteins for use in further applications. In polyacrylamide gel electrophoresis, proteins migrate in response to an electrical field through pores in a polyacrylamide gel matrix; pore size decreases with increasing acrylamide concentration. The combination of pore size and protein charge, size, and shape determines the migration rate of the protein. In this unit, the standard Laemmli method is described for discontinuous gel electrophoresis under denaturing conditions, i.e., in the presence of sodium dodecyl sulfate (SDS). 

In this video, we demonstrate a method for electrophoretic separation of proteins using poly-acrylimide gel electrophoresis (PAGE).

Electrophoresis is used to separate complex mixtures of proteins (e.g., from cells, subcellular fractions, column fractions, or immunoprecipitates), to ...

Harvesting Venom Toxins from Assassin Bugs and Other Heteropteran Insects

Heteropteran insects such as assassin bugs (Reduviidae) and giant water bugs (Belostomatidae) descended from a common predaceous and venomous ancestor, and the majority of extant heteropterans retain this trophic strategy. Some heteropterans have transitioned to feeding on vertebrate blood (such as the kissing bugs, Triatominae; and bed bugs, Cimicidae) while others have reverted to feeding on plants (most Pentatomomorpha). However, with the exception of saliva used by kissing bugs to facilitate blood-feeding, little is known about heteropteran venoms compared to the venoms of spiders, scorpions and snakes.
One obstacle to the characterization of heteropteran venom toxins is the structure and function of the venom/labial glands, which are both morphologically complex and perform multiple biological roles (defense, prey capture, and extra-oral digestion). In this article, we describe three methods we have successfully used to collect heteropteran venoms. First, we present electrostimulation as a convenient way to collect venom that is often lethal when injected into prey animals, and which obviates contamination by glandular tissue. Second, we show that gentle harassment of animals is sufficient to produce venom extrusion from the proboscis and/or venom spitting in some groups of heteropterans. Third, we describe methods to harvest venom toxins by dissection of anaesthetized animals to obtain the venom glands. This method is complementary to other methods, as it may allow harvesting of toxins from taxa in which electrostimulation and harassment are ineffective. These protocols will enable researchers to harvest toxins from heteropteran insects for structure-function characterization and possible applications in medicine and agriculture.

Although many insects in the suborder Heteroptera (Insecta: Hemiptera) are venomous, their venom composition and the functions of their venom toxins are mostly unknown. This protocol describes methods to harvest heteropteran venoms for further characterization, using electrostimulation, harassment, and gland dissection.

Heteropteran insects such as assassin bugs (Reduviidae) and giant water bugs (Belostomatidae) descended from a common predaceous and venomous ancestor, ...

Environment

Digital Microfluidics for Automated Proteomic Processing

Clinical proteomics has emerged as an important new discipline, promising the discovery of biomarkers that will be useful for early diagnosis and prognosis of disease. While clinical proteomic methods vary widely, a common characteristic is the need for (i) extraction of proteins from extremely heterogeneous fluids (i.e. serum, whole blood, etc.) and (ii) extensive biochemical processing prior to analysis. Here, we report a new digital microfluidics (DMF) based method integrating several processing steps used in clinical proteomics. This includes protein extraction, resolubilization, reduction, alkylation and enzymatic digestion. Digital microfluidics is a microscale fluid-handling technique in which nanoliter-microliter sized droplets are manipulated on an open surface. Droplets are positioned on top of an array of electrodes that are coated by a dielectric layer - when an electrical potential is applied to the droplet, charges accumulate on either side of the dielectric. The charges serve as electrostatic handles that can be used to control droplet position, and by biasing a sequence of electrodes in series, droplets can be made to dispense, move, merge, mix, and split on the surface. Therefore, DMF is a natural fit for carrying rapid, sequential, multistep, miniaturized automated biochemical assays. This represents a significant advance over conventional methods (relying on manual pipetting or robots), and has the potential to be a useful new tool in clinical proteomics.
Mais J. Jebrail, Vivienne N. Luk, and Steve C. C. Shih contributed equally to this work.
Sergio L. S. Freire's current address is at the University of the Sciences in Philadelphia located at 600 South 43rd Street, Philadelphia, PA 19104.

Digital Microfluidics is a technique characterized by the manipulation of discrete droplets (~nL - mL) on an array of electrodes by the application of electrical fields. It is well-suited for carrying out rapid, sequential, miniaturized automated biochemical assays. Here, we report a platform capable of automating several proteomic processing steps.

Clinical proteomics has emerged as an important new discipline, promising the discovery of biomarkers that will be useful for early diagnosis and ...

Toxin Induction and Protein Extraction from Fusarium spp. Cultures for Proteomic Studies

Fusaria are filamentous fungi able to produce different toxins. Fusarium mycotoxins such as deoxynivalenol, nivalenol, T2, zearelenone, fusaric acid, moniliformin, etc... have adverse effects on both human and animal health and some are considered as pathogenicity factors. Proteomic studies showed to be effective for deciphering toxin production mechanisms (Taylor et al., 2008) as well as for identifying potential pathogenic factors (Paper et al., 2007, Houterman et al., 2007) in Fusaria. It becomes therefore fundamental to establish reliable methods for comparing between proteomic studies in order to rely on true differences found in protein expression among experiments, strains and laboratories. The procedure that will be described should contribute to an increased level of standardization of proteomic procedures by two ways. The filmed protocol is used to increase the level of details that can be described precisely. Moreover, the availability of standardized procedures to process biological replicates should guarantee a higher robustness of data, taking into account also the human factor within the technical reproducibility of the extraction procedure.
The protocol described requires 16 days for its completion: fourteen days for cultures and two days for protein extraction (figure 1). 
Briefly, Fusarium strains are grown on solid media for 4 days; they are then manually fragmented and transferred into a modified toxin inducing media (Jiao et al., 2008) for 10 days. Mycelium is collected by filtration through a Miracloth layer. Grinding is performed in a cold chamber. Different operators performed extraction replicates (n=3) in order to take into account the bias due to technical variations (figure 2). Extraction was based on a SDS/DTT buffer as described in Taylor et al. (2008) with slight modifications. Total protein extraction required a precipitation process of the proteins using Aceton/TCA/DTT buffer overnight and Acetone /DTT washing (figure 3a,3b). Proteins were finally resolubilized in the protein-labelling buffer and quantified. Results of the extraction were visualized on a 1D gel (Figure 4, SDS-PAGE), before proceeding to 2D gels (IEF/SDS-PAGE). The same procedure can be applied for proteomic analyses on other growing media and other filamentous fungi (Miles et al., 2007).

Toxin Induction and Protein Extraction from Fusarium spp. Cultures for Proteomic Studies

Protein extraction for proteomic analyses in fungal species requires high levels of standardization to be accomplished according with the minimum information about a proteomic experiment (MIAPE) guidelines. We present a video-protocol that includes a procedure for minimizing experimental bias during toxin induction and protein extraction from Fusarium spp.

Fusaria are filamentous fungi able to produce different toxins. Fusarium mycotoxins such as deoxynivalenol, nivalenol, T2, zearelenone, fusaric acid, ...

Identification of Hemolytic and Phospholipase Activity in Crude Extracts from Sea Anemones by Straightforward Bioassays

Sea anemone venom composition includes polypeptide and non-proteins molecules. Cytolytic components have a high biotechnological and biomedical potential for designing new molecular tools. Sea anemone venom locates in glandular cells from ectoderm and sub-cellular structures called nematocysts, both of which are distributed throughout the sea anemone body. This characteristic implies challenges because the cells and nematocyst must be lysed to release the venom components with other non-toxic molecules. Therefore, first, the venom is derived from a crude extract (mixture of different and diverse molecules and tissue debris). The next step is to detect polypeptides with specific bioactivities. Here, we describe an efficient strategy to obtain the sea anemone crude extract and bioassay to identify the presence of cytolysins. The first step involves inexpensive and straightforward techniques (stirred and freeze-thaw cycle) to release cytolysins. We obtained the highest cytolytic activity and protein (~500 mg of protein from 20 g of dry weight). Next, the polypeptide complexity of the extract was analyzed by SDS-PAGE gel detecting proteins with molecular weights between 10 kDa and 250 kDa. In the hemolytic assay, we used sheep red blood cells and determined HU50 (11.1 &plusmn; 0.3 &#181;g/mL). In contrast, the presence of phospholipases in the crude extract was determined using egg yolk as a substrate in a solid medium with agarose. Overall, this study uses an efficient and inexpensive protocol to prepare the crude extract and applies replicable bioassays to identify cytolysins, molecules with biotechnological and biomedical interests.

Here, we describe a protocol to obtain crude venom extract from sea anemone and detect its hemolytic and phospholipase activity.

Sea anemone venom composition includes polypeptide and non-proteins molecules. Cytolytic components have a high biotechnological and biomedical potential ...

Biochemistry

Artificial Host Parasitization by Trichogramma dendrolimi and Venom Extraction

Extracting Venom from the Parasitoid Wasp Trichogramma dendrolimi Using an Artificial Host

Parasitoid wasps are a diverse group of hymenopteran insects that serve as invaluable resources for pest biocontrol. To ensure successful parasitism, parasitoid wasps inject venom into their hosts to suppress their hosts&#39; immunity, modulate hosts&#39; development, metabolism, and even behavior. With over 600,000 estimated species, the diversity of parasitoid wasps surpasses that of other venomous animals, such as snakes, cone snails, and spiders. Parasitoid wasp venom is an underexplored source of bioactive molecules with potential applications in pest control and medicine. However, collecting parasitoid venom is challenging due to the inability to use direct or electrical stimulation and the difficulty in dissection because of their small size. Trichogramma is a genus of tiny (~0.5 mm) egg parasitoid wasps that are widely used for the biological control of lepidopteran pests in both agriculture and forests. Here, we report a method for extracting venom from T. dendrolimi using artificial hosts. These artificial hosts are created with polyethylene film and amino acid solutions and then inoculated with Trichogramma wasps for parasitism. The venom was subsequently collected and concentrated. This method enables the extraction of large amounts of Trichogramma venom while avoiding contamination from other tissues caused by dissection, a common issue in venom reservoir dissection protocols. This innovative approach facilitates the study of Trichogramma venom, paving the way for new research and potential applications.

Author Spotlight: Efficient Venom Extraction Method from Trichogramma Parasitoid Wasps 

Here, we present a protocol for extracting venom from Trichogramma dendrolimi using an artificial host created with polyethylene film and amino acid solution.

Parasitoid wasps are a diverse group of hymenopteran insects that serve as invaluable resources for pest biocontrol. To ensure successful parasitism, ...

Microdissection of Black Widow Spider Silk-producing Glands

Modern spiders spin high-performance silk fibers with a broad range of biological functions, including locomotion, prey capture and protection of developing offspring 1,2. Spiders accomplish these tasks by spinning several distinct fiber types that have diverse mechanical properties. Such specialization of fiber types has occurred through the evolution of different silk-producing glands, which function as small biofactories. These biofactories manufacture and store large quantities of silk proteins for fiber production. Through a complex series of biochemical events, these silk proteins are converted from a liquid into a solid material upon extrusion.
Mechanical studies have demonstrated that spider silks are stronger than high-tensile steel 3. Analyses to understand the relationship between the structure and function of spider silk threads have revealed that spider silk consists largely of proteins, or fibroins, that have block repeats within their protein sequences 4. Common molecular signatures that contribute to the incredible tensile strength and extensibility of spider silks are being unraveled through the analyses of translated silk cDNAs. Given the extraordinary material properties of spider silks, research labs across the globe are racing to understand and mimic the spinning process to produce synthetic silk fibers for commercial, military and industrial applications. One of the main challenges to spinning artificial spider silk in the research lab involves a complete understanding of the biochemical processes that occur during extrusion of the fibers from the silk-producing glands.
Here we present a method for the isolation of the seven different silk-producing glands from the cobweaving black widow spider, which includes the major and minor ampullate glands [manufactures dragline and scaffolding silk] 5,6, tubuliform [synthesizes egg case silk] 7,8, flagelliform [unknown function in cob-weavers], aggregate [makes glue silk], aciniform [synthesizes prey wrapping and egg case threads] 9 and pyriform [produces attachment disc silk] 10. This approach is based upon anesthetizing the spider with carbon dioxide gas, subsequent separation of the cephalothorax from the abdomen, and microdissection of the abdomen to obtain the silk-producing glands. Following the separation of the different silk-producing glands, these tissues can be used to retrieve different macromolecules for distinct biochemical analyses, including quantitative real-time PCR, northern- and western blotting, mass spectrometry (MS or MS/MS) analyses to identify new silk protein sequences, search for proteins that participate in the silk assembly pathway, or use the intact tissue for cell culture or histological experiments.

Here we describe an efficient strategy to remove the silk-producing glands from the abdomen of female black widow spiders. This procedure allows the rapid isolation of the seven distinct silk-producing glands in a highly purified fashion, an important process for investigators studying spider silk production and fiber assembly.

Modern spiders spin high-performance silk fibers with a broad range of biological functions, including locomotion, prey capture and protection of ...

High Throughput Quantitative Expression Screening and Purification Applied to Recombinant Disulfide-rich Venom Proteins Produced in E. coli

Escherichia coli (E. coli) is the most widely used expression system for the production of recombinant proteins for structural and functional studies. However, purifying proteins is sometimes challenging since many proteins are expressed in an insoluble form. When working with difficult or multiple targets it is therefore recommended to use high throughput (HTP) protein expression screening on a small scale (1-4&#160;ml cultures) to quickly identify conditions for soluble expression. To cope with the various structural genomics programs of the lab, a quantitative (within a range of 0.1-100 mg/L culture of recombinant protein) and HTP protein expression screening protocol was implemented and validated on thousands of proteins. The protocols were automated with the use of a liquid handling robot but can also be performed manually without specialized equipment.
Disulfide-rich venom proteins are gaining increasing recognition for their potential as therapeutic drug leads. They can be highly potent and selective, but their complex disulfide bond networks make them challenging to produce. As a member of the FP7 European Venomics project (<a href="http://www.venomics.eu">www.venomics.eu</a>), our challenge is to develop successful production strategies with the aim of producing thousands of novel venom proteins for functional characterization. Aided by the redox properties of disulfide bond isomerase DsbC, we adapted our HTP production pipeline for the expression of oxidized, functional venom peptides in the E. coli cytoplasm. The protocols are also applicable to the production of diverse disulfide-rich proteins. Here we demonstrate our pipeline applied to the production of animal venom proteins. With the protocols described herein it is likely that soluble disulfide-rich proteins will be obtained in as little as a week. Even from a small scale, there is the potential to use the purified proteins for validating the oxidation state by mass spectrometry, for characterization in pilot studies, or for sensitive micro-assays.

High Throughput Quantitative Expression Screening and Purification Applied to Recombinant Disulfide-rich Venom Proteins Produced in E. coli

A protocol for the quantitative, high throughput expression screening and analytical purification of fusion proteins from small-scale Escherichia coli cultures is described and applied to the expression of disulfide-rich animal venom protein targets.

Escherichia coli (E. coli) is the most widely used expression system for the production of recombinant proteins for structural and functional studies. ...

Rearing of Brazilian Yellow Scorpion for Venom Extraction

Captive Maintenance and Venom Extraction of Tityus serrulatus (Brazilian Yellow Scorpion) for Antivenom Production

Scorpion envenomation is a public health problem in several tropical and subtropical countries. Tityus serrulatus Lutz and Mello, 1922 (Brazilian yellow scorpion) are responsible for approximately 150,000 envenoming cases per year in Brazil, of which 10% require antivenom treatment to reverse life-threatening venom effects. Therefore, thousands of T. serrulatus individuals are maintained under controlled captivity conditions for venom extraction, subsequently used in the production of the national supply of scorpion antivenom. Instituto Butantan is the main antivenom-manufacturing laboratory in Brazil, providing about 70,000 vials of scorpion antivenom for the Brazilian health system. Thus, the husbandry protocols and venom extraction methodologies are key points for the success of large-scale, standardized venom production. The objective of this article is to describe the captivity protocols of T. serrulatus husbandry, encompassing the husbandry routine and the venom extraction procedures, following good manufacturing practices, and ensuring animal welfare. These practices allow for the maintenance of up to 20,000 animals in captivity, with a routine of 3,000 to 5,000 scorpions milked monthly according to antivenom manufacturing demand, achieving an average of 90% of positive extraction.

Author Spotlight: Optimizing Scorpion Venom Extraction for Antivenom Production 

Here, we present successful maintenance and venom extraction protocols for large-scale Tityus serrulatus Lutz and Mello, 1922 (Brazilian yellow scorpion) husbandry, with the aim of providing venom for subsequent scorpion antivenom production to meet the demand of the Brazilian health system.

Scorpion envenomation is a public health problem in several tropical and subtropical countries. Tityus serrulatus Lutz and Mello, 1922 (Brazilian yellow ...

Phospholipase Activity Assay: A Gel Diffusion Assay to Determine Phospholipase Activity in Crude Venom Extract from Anthopleura dowii

Source: Ram&#237;rez-Carreto, S., et al., <a href="https://www-jove-com.remotexs.ntu.edu.sg/v/63630">Identification of Hemolytic and Phospholipase Activity in Crude Extracts from Sea Anemones by Straightforward Bioassays.</a> J. Vis. Exp. (2022)
In this video, we demonstrate an assay to establish the presence of phospholipase in the crude venom extract from Anthopleura dowii&#8212;a marine sea anemone. The assay utilizes an agarose gel diffusion technique, in which the enzyme phospholipase diffuses and breaks down phospholipids incorporated in the solid media in a radial manner.

Source: Ram&#237;rez-Carreto, S., et al., Identification of Hemolytic and Phospholipase Activity in Crude Extracts from Sea Anemones by Straightforward ...

Extraction of Venom and Venom Gland Microdissections from Spiders for Proteomic and Transcriptomic Analyses

Summary

Explore More Videos

Chapters in this video

Electrophoretic Separation of Proteins

Harvesting Venom Toxins from Assassin Bugs and Other Heteropteran Insects

Digital Microfluidics for Automated Proteomic Processing

Toxin Induction and Protein Extraction from Fusarium spp. Cultures for Proteomic Studies

Identification of Hemolytic and Phospholipase Activity in Crude Extracts from Sea Anemones by Straightforward Bioassays

Extracting Venom from the Parasitoid Wasp Trichogramma dendrolimi Using an Artificial Host

Microdissection of Black Widow Spider Silk-producing Glands

High Throughput Quantitative Expression Screening and Purification Applied to Recombinant Disulfide-rich Venom Proteins Produced in E. coli

Captive Maintenance and Venom Extraction of Tityus serrulatus (Brazilian Yellow Scorpion) for Antivenom Production

Phospholipase Activity Assay: A Gel Diffusion Assay to Determine Phospholipase Activity in Crude Venom Extract from Anthopleura dowii