We thank William T. Eckert for narrating the video. Development of the glass wool protocol was part of the Wisconsin Water And Health Trial for Enteric Risks (WAHTER Study), funded by US EPA STAR Grant R831630. Alaska samples were collected by A. Reeves, A. Ramey, and B. Meixell with financial support from USGS. Any use of trade, product, or firm names is for descriptive purposes only and does not imply endorsement by the U.S. Government.

1. Preparing the Glass Wool
<ol>
 <li> Before and after making each batch of filters, sterilize the work area with 10% bleach solution. </li>
 <li> Put on gloves and gown. Sterilize a bucket by autoclaving at 121&deg;C and 15 psi for at least 20 minutes. Place the glass wool in the sterile bucket. </li>
 <li> Saturate the glass wool with reverse osmosis water and let soak for 15 minutes. </li>
 <li> Drain the reverse osmosis water from the bucket.</li>
 <li> Saturate the glass wool with 1 M HCl and let soak for 15 minutes. </li>
 <li> Drain the 1 M HCl from the bucket.</li>
 <li> Rinse the glass wool with reverse osmosis water. </li>
 <li> Mix thoroughly. </li>
 <li> Check the pH using pH paper and repeat the reverse osmosis water rinse until a neutral pH is achieved. </li>
 <li> Pour off the rinse water. </li>
 <li> Saturate the glass wool with 1 M NaOH and let soak for 15 minutes. </li>
 <li> Drain the 1 M NaOH from the bucket. </li>
 <li> Repeat the reverse osmosis rinse until a neutral pH is achieved. </li>
 <li> Pour off the rinse water. </li>
 <li> Instead of a bucket, a glass wool washer can be constructed, which is similar in design to a glass pipette washer (Figure 1). </li>
 <li> Cover the glass wool completely with sterile Phosphate Buffered Saline (PBS) adjusted to pH 6.8. </li>
 <li> Use prepared glass wool immediately or store at 4&deg;C. It can be stored for up to two weeks. Before using, make sure the pH is neutral as it will rise over time. If the pH is not neutral, re-rinse with sterile Phosphate Buffered Saline (pH 6.8). </li>
</ol>

2. Assembling the Glass Wool Filter
<ol>
 <li> Drill a 11/16 inch hole into the PVC caps and tap threads so the male adapter nylon fittings can be screwed into the caps. This step is necessary only for the first assembly. Thereafter, the caps can be used for years. Apply Teflon tape to the nylon fittings threads and screw to the caps. </li>
 <li> Pack the PVC pipe with small pieces of glass wool. Use a metal plunger, like a car engine valve, to pack tight. Packing does not require great force. Pack tight enough so that the glass wool stays in place and channels do not form. However, do not pack so tightly water cannot flow through the filter. When packed appropriately, a flow rate of 4 to 5 liters per minute should be attainable when the filter is attached to water taps with pressures between 40-60 psi. For the size of PVC pipe specified in this protocol, approximately 85 grams washed and packed glass wool is used per pipe. Tare the empty PVC pipe on a top-loading balance and pack with washed glass wool until the pipe mass increases 85 grams. </li>
 <li> Insert polypropylene mesh into the PVC caps with male adapter nylon fittings attached.</li>
 <li> Apply Teflon tape to the threaded portions of the PVC pipe. </li>
 <li> Screw on PVC caps to the PVC pipe and label one filter end the inflow and the other end the outflow. This is important later for the elution step. Which end is labeled inflow does not matter. </li>
 <li> Push 60 mL of sterile Phosphate Buffered Saline (pH = 6.8) into the filter using a catheter tipped syringe. Excess will come out opposite end. </li>
 <li> Wrap ends tightly with Parafilm to avoid leaking. Filters can be stored up to 30 days at 4&deg;C. </li>
</ol>
3. Sampling
<ol>
 <li> Sterilize tubing used for sampling by recirculating or immersing items for 30 minutes in 0.525% NaClO (i.e., a 10% solution of standard household bleach). Follow this by draining the hypochlorite solution and neutralize with 0.05% sodium thiosulfate, made by adding 25 mL of a 2% stock anhydrous sodium thiosulfate solution to one liter of reverse osmosis water. </li>
 <li> For sample water with a pH greater than 7.5, adjust the pH to between 6.5 and 7.0 with HCl. HCl concentration can range from 0.25 M to 1 M. Four liters 0.5 M HCl is generally sufficient for two 800 liter samples, depending on the water's ambient pH and buffering capacity. Inject HCl during sampling using a peristaltic pump or Venturi (Figure 2). Adjust the pump speed or Venturi opening to achieve the target pH. Measure the pH at the sample line outlet using a field pH meter.</li>
 <li> Use a prefilter if the glass wool filter clogs from water with high turbidity. (Specifications for the prefilter and its housing are in the Materials list online.) Because pathogens can be attached to particles trapped by the prefilter it must be eluted along with the glass wool filter (see below). </li>
 <li> Adjust the flow rate to between 2 and 4 liters/minute. Typical sample volumes are between 200 and 1,500 liters. </li>
 <li> Disconnect the glass wool filter and store in a plastic sterile bag at 4&deg;C for up to 48 hours. </li>
</ol>
4. Elution
<ol>
 <li> Affix the glass wool filter to a ring stand with the sample inflow end pointing downward into a polypropylene bottle. Elute opposite of the direction of sample flow. </li>
 <li> Push 80 mL of sterile 3% beef extract in 0.05 M glycine with a pH of 9.5 into the filter.</li>
 <li> Wait 15 minutes.</li>
 <li> Push another 80 mL of sterile 3% beef extract in 0.05 M glycine with a pH of 9.5 into the filter. </li>
 <li> Push air thorough the filter until foam comes out of the filter inlet. </li>
 <li> Adjust the eluate pH to between 7.0 and 7.5 with 1 M HCl. </li>
 <li> Store eluate at 4&deg;C for up to 24 hours or at -20&deg;C for longer periods of time. </li>
 <li> Elute the prefilter if one is used. Unscrew the top of the housing cartridge and pour off all residual water inside the housing while holding the prefilter in place with sterile gloved hands. </li>
 <li> Remove the prefilter from the housing cartridge and slip it into a 15" x 6" zip-lock bag. </li>
 <li> Pour 200 mL of sterile 3% beef extract in 0.05 M glycine with a pH of 9.5 into the bag and seal securely with the zip-lock. </li>
 <li> Invert the bagged prefilter several times to ensure the entire surface comes in contact with the beef extract.</li>
 <li> Wait 15 minutes, occasionally inverting and massaging the bagged prefilter.</li>
 <li> Open the zip-lock. Grip the bag around the prefilter tightly to squeeze out as much beef extract as possible and pull the prefilter out with sterile gloved hands,. </li>
 <li> Pour the beef extract eluate into a polypropylene bottle.</li>
 <li> Adjust the eluate pH to between 7.0 and 7.5 with 1M HCl. </li>
 <li> Store eluate at 4&deg;C for up to 24 hours or at -20&deg;C for longer periods of time. The prefilter eluate can be separately analyzed for pathogens or combined with the glass wool filter eluate. </li>
</ol>









5. Representative Results 
<table border="1">
<tbody>
 <tr>
 <td>Pathogen</td>
 <td>Water Turbidity Level (NTU) a</td>
 <td>Amount Seeded/L&nbsp;b,c,d</td>
 <td>% Recovery &plusmn; 1 SD</td>
 <td>Number Independent 
Trials</td>
 </tr>
 <tr>
 <td>Enterovirus- Poliovirus Sabin III</td>
 <td>0.5 </td>
 <td>500</td>
 <td>81% &plusmn; 11</td>
 <td>7</td>
 </tr>
 <tr>
 <td>Enterovirus- Poliovirus Sabin III</td>
 <td>0.5</td>
 <td>5000</td>
 <td>67% &plusmn; 12</td>
 <td>8</td>
 </tr>
 <tr>
 <td>Enterovirus- Poliovirus Sabin III</td>
 <td>215 </td>
 <td>500</td>
 <td>59% &plusmn; 32</td>
 <td>7</td>
 </tr>
 <tr>
 <td>Enterovirus- Poliovirus Sabin III</td>
 <td>215</td>
 <td>5000</td>
 <td>38% &plusmn; 22</td>
 <td>6</td>
 </tr>
 <tr>
 <td>Enterovirus- Poliovirus Sabin III</td>
 <td>447</td>
 <td>500</td>
 <td>56% &plusmn; 18</td>
 <td>8</td>
 </tr>
 <tr>
 <td>Enterovirus- Poliovirus Sabin III</td>
 <td>447</td>
 <td>5000</td>
 <td>63% &plusmn; 37</td>
 <td>8</td>
 </tr>
 <tr>
 <td>Cryptosporidium parvum</td>
 <td>0.5</td>
 <td>5</td>
 <td>38% &plusmn; 14</td>
 <td>7</td>
 </tr>
 <tr>
 <td>Cryptosporidium parvum</td>
 <td>0.5</td>
 <td>50</td>
 <td>53% &plusmn; 19</td>
 <td>8</td>
 </tr>
 <tr>
 <td>Cryptosporidium parvum</td>
 <td>215</td>
 <td>5</td>
 <td>40% &plusmn; 16</td>
 <td>7</td>
 </tr>
 <tr>
 <td>Cryptosporidium parvum</td>
 <td>215</td>
 <td>50</td>
 <td>30% &plusmn;6</td>
 <td>6</td>
 </tr>
 <tr>
 <td>Cryptosporidium parvum</td>
 <td>447</td>
 <td>5</td>
 <td>33% &plusmn; 13</td>
 <td>8</td>
 </tr>
 <tr>
 <td>Cryptosporidium parvum</td>
 <td>447</td>
 <td>50</td>
 <td>28% &plusmn; 11</td>
 <td>8</td>
 </tr>
 <tr>
 <td>Salmonella enterica</td>
 <td>0.5</td>
 <td>5</td>
 <td>29% &plusmn; 24</td>
 <td>7</td>
 </tr>
 <tr>
 <td>Salmonella enterica</td>
 <td>0.5</td>
 <td>500</td>
 <td>56% &plusmn; 16</td>
 <td>8</td>
 </tr>
 <tr>
 <td>Salmonella enterica</td>
 <td>215</td>
 <td>5</td>
 <td>32% &plusmn; 24</td>
 <td>7</td>
 </tr>
 <tr>
 <td>Salmonella enterica</td>
 <td>215</td>
 <td>500</td>
 <td>34% &plusmn;11</td>
 <td>6</td>
 </tr>
 <tr>
 <td>Salmonella enterica</td>
 <td>447</td>
 <td>5</td>
 <td>34% &plusmn; 18</td>
 <td>8</td>
 </tr>
 <tr>
 <td>Salmonella enterica</td>
 <td>447</td>
 <td>500</td>
 <td>31% &plusmn; 24</td>
 <td>8</td>
 </tr>
 </tbody>
</table>
<ol class="lalpha">
<li> Nephelometric Turbidity Unit</li>
<li> Enteroviruses enumerated by qPCR as genomic copies/L</li>
<li> C. parvum enumerated by immunofluorescence as oocysts</li>
<li> S. enterica enumerated by culture as colony-forming-units</li>
</ol>
Table 1. Glass wool concentration with varying water turbidities and pathogen densities.
 <table border="1">
 <tbody>
 <tr>
 <td>Pathogen</td>
 <td>Water Sample Location</td>
 <td>Amount Seeded/La</td>
 <td>% Recovery</td>
 </tr>
 <tr>
 <td>Avian Influenza H5N2</td>
 <td>Sundi Lake, Anchorage Borough</td>
 <td>2500</td>
 <td>42.9%</td>
 </tr>
 <tr>
 <td>Avian Influenza H5N2</td>
 <td>Minto Flats, Fairbanks North Star Borough</td>
 <td>2500</td>
 <td>36.7%</td>
 </tr>
 <tr>
 <td>Avian Influenza H5N2</td>
 <td>Portage Valley, Anchorage Borough</td>
 <td>2500</td>
 <td>7.8%</td>
 </tr>
 <tr>
 <td>Avian Influenza H5N2</td>
 <td>Potter Marsh, Anchorage Borough</td>
 <td>2500</td>
 <td>41.5%</td>
 </tr>
 <tr>
 <td>Avian Influenza H5N2</td>
 <td>Willow Lake, Yukon-Koyukuk Borough</td>
 <td>2500</td>
 <td>15.5%</td>
 </tr>
 </tbody>
 </table>
<ol class="lalpha">
<li> Measured by qPCR as genomic copies/L</li>
</ol>
Table 2. Glass wool concentration of avian influenza virus using water from five sites in Alaska.
<img src="/files/ftp_upload/3930/3930fig1.jpg" alt="Figure 1" /> 
Figure 1. Diagram of glass wool washer. This can be used instead of a bucket, saving time from rinsing. The concept is similar to a glass pipette washer.
<img src="/files/ftp_upload/3930/3930fig2.jpg" alt="Figure 2" /> 
Figure 2. Glass wool filtration with acid injection by peristaltic pump. Note the "T" connector where the pump tubing introduces acid into the sample line between the water tap and glass wool filter. pH adjustment is necessary only if the water sampled has a pH &gt; 7.5. 
Glass wool filters are effective in concentrating pathogens from water with a wide range of turbidity levels and pathogen densities (Table 1). To test this, 20 liters dechlorinated tap water was mixed with dried silt loam soil (0, 1.27, or 2.75 g/L) to achieve the desired level of turbidity and then seeded with pathogens at various densities. The water was passed through a glass wool filter, the concentrated pathogens in the eluate were enumerated, and this value was the numerator in the percent recovery calculation. The quantity of pathogens seeded into the water, that is, the denominator of the percent recovery calculation, was determined by first seeding the pathogens into a negative eluate then enumerating the pathogens. The negative eluate was prepared by passing an unseeded 20 liter sample through a filter and eluting. Quantifying the seeded pathogens in a negative eluate avoids differences in pathogen enumeration that could result from matrix differences created by the glass wool filter. The importance of this step when quantifying pathogens by qPCR is discussed in Lambertini et al.3. A 20 liter negative control sample was concentrated via glass wool filtration to determine there were no native pathogens present that could confound the percent recovery calculation. A 10 &mu;m nominal pore size prefilter was used when the turbidity level was &ge; 215 NTU.
Poliovirus was quantified by real-time qPCR using the secondary concentration and nucleic acid extraction procedures and primers and probe described in Lambertini et al.3. Cryptosporidium parvum was quantified in the final concentrated sample volume (FCSV) created by the secondary concentration procedure for poliovirus. Oocysts were visualized by immunofluorescence (MeriFluor Cryptosporidium &amp; Giardia Detection Kit, Meridian Life Science, Inc., Cincinnati, OH). Salmonella enterica was quantified in the FCSV by plating onto XLD agar (Remel, Lenexa, KS) and counting colony-forming-units.
Glass wool filters are effective in concentrating avian influenza viruses (Table 2). Low pathogenicity avian influenza virus (H5N2) was seeded into water from several locations in Alaska and the percent recovery calculated as described above. Secondary concentration and nucleic acid procedures were performed as for poliovirus; the virus was quantified by qPCR using the primers and probe described in Spackman et al.4

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	<li>US Environmental Protection Agency. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Method+1623:+Cryptosporidium+and+Giardia+in+Water+by+Filtration/IMS/FA.">Method 1623: Cryptosporidium and Giardia in Water by Filtration/IMS/FA.</a> EPA 815-R-05-002. , (2012).</li><li>Cashdollar, J. L., Dahling, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+a+method+to+re-use+electropositive+cartridge+filters+for+concentrating+viruses+from+tap+and+river+water.">Evaluation of a method to re-use electropositive cartridge filters for concentrating viruses from tap and river water.</a> J. Virol. Methods. 132, 13-17 (2006).</li><li>Lambertini, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Concentration+of+enteroviruses,+adenoviruses,+and+noroviruses+from+drinking+water+by+use+of+glass+wool+filters.">Concentration of enteroviruses, adenoviruses, and noroviruses from drinking water by use of glass wool filters.</a> Appl. Environ. Microbiol. 74, 2990-2996 (2008).</li><li>Spackman, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+real-time+reverse+transcription+PCR+assay+for+Type+A+influenza+virus+and+the+avian+H5+and+H7+hemagglutinin+subtypes.">Development of a real-time reverse transcription PCR assay for Type A influenza virus and the avian H5 and H7 hemagglutinin subtypes.</a> J. Clin. 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No conflicts of interests declared.

Glass wool filters have been used by several research teams3,5,6 to concentrate human enteric viruses from a variety of water sources such as finished drinking water7, groundwater8,9, surface water10, sea water11, wastewater12, and agricultural runoff13. Here we report the filters are also effective in concentrating avian influenza virus as well as the bacterial and protozoan pathogens Salmonella enterica (serovar Typhimurium) and C...

<table border="1">
<tbody>
 <tr>
 <td>Name of reagent or item</td>
 <td> Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Hydrochloric acid</td>
 <td><a href="http://www.fishersci.com" target="_blank">Fisher Scientific</a></td>
 <td>A144-500</td>
 </tr>
 <tr>
 <td height="24">Sodium hydroxide</td>
 <td><a href="http://www.fishersci.com" target="_blank">Fisher Scientific</a></td>
 <td>BP359-212</td>
 </tr>
 <tr>
 <td>Phosphate Buffered Saline 
 Sodium chloride
 
 Potassium phosphate-dibasic
 
 Potassium phosphate-monobasic</td>
 <td>
 
 <a href="http://www.fishersci.com" target="_blank">Fisher Scientific</a> 
 <a href="http://www.fishersci.com" target="_blank">Fisher Scientific</a> 
 <a href="http://www.fishersci.com" target="_blank">Fisher Scientific</a></td>
 <td>
 
 BP358-212
 
 BP363-500
 
 BP362-500</td>
 </tr>
 <tr>
 <td>Sodium hypochlorite
 i.e., household bleach</td>
 <td><a href="http://www.clorox.com" target="_blank">The Clorox Co.</a></td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Sodium thiosulfate, anhydrous</td>
 <td><a href="http://www.fishersci.com" target="_blank">Fisher Scientific</a></td>
 <td>S 475-212</td>
 </tr>
 <tr>
 <td>Beef extract, desiccated</td>
 <td> <a href="http://www.bd.com" target="_blank">Becton, Dickinson and Company</a></td>
 <td>211520</td>
 </tr>
 <tr>
 <td>Glycine</td>
 <td><a href="http://www.fishersci.com" target="_blank">Fisher Scientific</a></td>
 <td>G46-500</td>
 </tr>
 <tr>
 <td>Oiled sodocalcic glass wool
 
 Or
 
 R-11 unfaced fiberglass insulation </td>
 <td> <a href="http://www.isover.com" target="_blank">Isover</a>
 
 <a href="http://www.jm.com" target="_blank">Johns Manville</a></td>
 <td>Bourre 725 QN </td>
 </tr>
 <tr>
 <td>Polypropylene mesh</td>
 <td> <a href="http://www.industrialnetting.com" target="_blank">Industrial Netting</a></td>
 <td>xN4510</td>
 </tr>
 <tr>
 <td>2"x4" Sch 80 PVC threaded pipe nipple</td>
 <td> <a href="http://www.grainger.com" target="_blank">Grainger</a></td>
 <td>6MW35</td>
 </tr>
 <tr>
 <td>2" Sch 40 PVC cap</td>
 <td><a href="http://www.grainger.com" target="_blank">Grainger</a></td>
 <td>5WDW3</td>
 </tr>
 <tr>
 <td>Male adapter nylon fitting (1/2"x1/2")</td>
 <td><a href="http://www.usplastic.com" target="_blank"> US Plastic Corp.</a></td>
 <td>62178</td>
 </tr>
 <tr>
 <td>Sample bottles for eluate- 1 liter</td>
 <td><a href="http://www.fishersci.com" target="_blank">Fisher Scientific</a></td>
 <td>03-313-4F</td>
 </tr>
 <tr>
 <td>60 mL syringe</td>
 <td><a href="http://www.fishersci.com" target="_blank">Fisher Scientific</a></td>
 <td>NC9661991</td>
 </tr>
 <tr>
 <td>pH strips</td>
 <td><a href="http://www.whatman.com" target="_blank">Whatman</a></td>
 <td>2614 991</td>
 </tr>
 <tr>
 <td>Prefilter, Polypropylene, 10 inch cartridge, 10 &mu;m</td>
 <td>McMaster-Carr</td>
 <td>4411K75</td>
 </tr>
 <tr>
 <td>Prefilter housing</td>
 <td>Cole-Parmer</td>
 <td>S-29820-10</td>
 </tr>
</tbody>
</table>

glass wool filters for concentrating waterborne viruses and agricultural zoonotic pathogens

The key first step in evaluating pathogen levels in suspected contaminated water is concentration. Concentration methods tend to be specific for a particular pathogen group, for example US Environmental Protection Agency Method 1623 for Giardia and Cryptosporidium1, which means multiple methods are required if the sampling program is targeting more than one pathogen group. Another drawback of current methods is the equipment can be complicated and expensive, for example the VIRADEL method with the 1MDS cartridge filter for concentrating viruses2. In this article we describe how to construct glass wool filters for concentrating waterborne pathogens. After filter elution, the concentrate is amenable to a second concentration step, such as centrifugation, followed by pathogen detection and enumeration by cultural or molecular methods. The filters have several advantages. Construction is easy and the filters can be built to any size for meeting specific sampling requirements. The filter parts are inexpensive, making it possible to collect a large number of samples without severely impacting a project budget. Large sample volumes (100s to 1,000s L) can be concentrated depending on the rate of clogging from sample turbidity. The filters are highly portable and with minimal equipment, such as a pump and flow meter, they can be implemented in the field for sampling finished drinking water, surface water, groundwater, and agricultural runoff. Lastly, glass wool filtration is effective for concentrating a variety of pathogen types so only one method is necessary. Here we report on filter effectiveness in concentrating waterborne human enterovirus, Salmonella enterica, Cryptosporidium parvum, and avian influenza virus.

Watch this Scientific Journal Video about Glass Wool Filters for Concentrating Waterborne Viruses and Agricultural Zoonotic Pathogens at JoVE.com

Glass Wool Filters for Concentrating Waterborne Viruses and Agricultural Zoonotic Pathogens

Glass wool filters have been used to concentrate waterborne viruses by a number of research groups around the world. Here we show a simple approach for constructing glass wool filters and demonstrate the filters are also effective in concentrating waterborne viral, bacterial and protozoan pathogens.

The key first step in evaluating pathogen levels in suspected contaminated water is concentration.  Concentration methods tend to be specific for a ...

glass-wool-filters-for-concentrating-waterborne-viruses-agricultural

Research

JoVE Journal

Immunology and Infection

17.9K ビュー.  United States Geological Survey. グラスウールフィルタは、世界中の研究グループの数によって水系ウイルスを濃縮するために使用されている。ここでは、グラスウールフィルタを構築するための単純なアプローチを示し、フィルタはまた、水性、ウイルス、細菌や原生動物病原体を濃縮するのに有効であるを示しています。

ビデオ: Glass Wool Filters for Concentrating Waterborne Viruses and Agricultural Zoonotic Pathogens

Methods for Rapid Transfer and Localization of Lyme Disease Pathogens Within the Tick Gut

Lyme disease is caused by infection with the spirochete pathogen Borrelia burgdorferi, which is maintained in nature by a tick-rodent infection cycle 1. A tick-borne murine model 2 has been developed to study Lyme disease in the laboratory. While na&iacute;ve ticks can be infected with B. burgdorferi by feeding them on infected mice, the molting process takes several weeks to months to complete. Therefore, development of more rapid and efficient tick infection techniques, such as a microinjection-based procedure, is an important tool for the study of Lyme disease 3,4. The procedure requires only hours to generate infected ticks and allows control over the delivery of equal quantities of spirochetes in a cohort of ticks. This is particularly important as the generation of B. burgdorferi infected ticks by the natural feeding process using mice fails to ensure 100% infection rate and potentially results in variation of pathogen burden amongst fed ticks. Furthermore, microinjection can be used to infect ticks with B. burgdorferi isolates in cases where an attenuated strain is unable to establish infection in mice and thus can not be naturally acquired by ticks 5. This technique can also be used to deliver a variety of other biological materials into ticks, for example, specific antibodies or double stranded RNA 6. In this article, we will demonstrate the microinjection of nymphal ticks with in vitro-grown B. burgdorferi. We will also describe a method for localization of Lyme disease pathogens in the tick gut using confocal immunofluorescence microscopy.

Lyme disease research studies often require generation of ticks infected with the pathogen Borrelia burgdorferi, a process that typically takes several weeks. Here we demonstrate a microinjection-based tick infection procedure that can be accomplished within hours. We also demonstrate an immunofluorescence method for in situ localization of B. burgdorferi within ticks.

Lyme disease is caused by infection with the spirochete pathogen Borrelia burgdorferi, which is maintained in nature by a tick-rodent infection cycle 1.  ...

'Bioluminescent' Reporter Phage for the Detection of Category A Bacterial Pathogens

Yersinia pestis and Bacillus anthracis are Category A bacterial pathogens that are the causative agents of the plague and anthrax, respectively 1. Although the natural occurrence of both diseases&#39; is now relatively rare, the possibility of terrorist groups using these pathogens as a bioweapon is real. Because of the disease&#39;s inherent communicability, rapid clinical course, and high mortality rate, it is critical that an outbreak be detected quickly. Therefore methodologies that provide rapid detection and diagnosis are essential to ensure immediate implementation of public health measures and activation of crisis management.
Recombinant reporter phage may provide a rapid and specific approach for the detection of Y. pestis and B. anthracis. The Centers for Disease Control and Prevention currently use the classical phage lysis assays for the confirmed identification of these bacterial pathogens 2-4. These assays take advantage of naturally occurring phage which are specific and lytic for their bacterial hosts. After overnight growth of the cultivated bacterium in the presence of the specific phage, the formation of plaques (bacterial lysis) provides a positive identification of the bacterial target. Although these assays are robust, they suffer from three shortcomings: 1) they are laboratory based; 2) they require bacterial isolation and cultivation from the suspected sample, and 3) they take 24-36 h to complete. To address these issues, recombinant &#34;light-tagged&#34; reporter phage were genetically engineered by integrating the Vibrio harveyi luxAB genes into the genome of Y. pestis and B. anthracis specific phage 5-8. The resulting luxAB reporter phage were able to detect their specific target by rapidly (within minutes) and sensitively conferring a bioluminescent phenotype to recipient cells. Importantly, detection was obtained either with cultivated recipient cells or with mock-infected clinical specimens 7.
For demonstration purposes, here we describe the method for the phage-mediated detection of a known Y. pestis isolate using a luxAB reporter phage constructed from the CDC plague diagnostic phage &#934;A1122 6,7 (Figure 1). A similar method, with minor modifications (e.g. change in growth temperature and media), may be used for the detection of B. anthracis isolates using the B. anthracis reporter phage W&#946;::luxAB 8. The method describes the phage-mediated transduction of a biolumescent phenotype to cultivated Y. pestis cells which are subsequently measured using a microplate luminometer. The major advantages of this method over the traditional phage lysis assays is the ease of use, the rapid results, and the ability to test multiple samples simultaneously in a 96-well microtiter plate format.
<img alt="Figure 1" src="/files/ftp_upload/2740/2740fig1.jpg" /> 
Figure 1. Detection schematic. The phage are mixed with the sample, the phage infects the cell, luxAB are expressed, and the cell bioluminesces. Sample processing is not necessary; the phage and cells are mixed and subsequently measured for light.

&#039;Bioluminescent&#039; Reporter Phage for the Detection of Category A Bacterial Pathogens

A simple method for the identification of priority bacterial pathogens is to use genetically engineered reporter phage. These reporter phage, which are specific to their particular host species, are capable of rapidly transducing a bioluminescent signal response to host cells. Herein, we describe the use of reporter phage for the detection of Yersinia pestis.

Yersinia pestis and Bacillus anthracis are Category A bacterial pathogens that are the causative agents of the plague and anthrax, respectively 1. ...

Avian Influenza Surveillance with FTA  Cards: Field Methods, Biosafety, and Transportation Issues Solved

Avian Influenza Viruses (AIVs) infect many mammals, including humans1. These AIVs are diverse in their natural hosts, harboring almost all possible viral subtypes2. Human pandemics of flu originally stem from AIVs3. Many fatal human cases during the H5N1 outbreaks in recent years were reported. Lately, a new AIV related strain swept through the human population, causing the 'swine flu epidemic'4. Although human trading and transportation activity seems to be responsible for the spread of highly pathogenic strains5, dispersal can also partly be attributed to wild birds6, 7. However, the actual reservoir of all AIV strains is wild birds.
In reaction to this and in face of severe commercial losses in the poultry industry, large surveillance programs have been implemented globally to collect information on the ecology of AIVs, and to install early warning systems to detect certain highly pathogenic strains8-12. Traditional virological methods require viruses to be intact and cultivated before analysis. This necessitates strict cold chains with deep freezers and heavy biosafety procedures to be in place during transport. Long-term surveillance is therefore usually restricted to a few field stations close to well equipped laboratories. Remote areas cannot be sampled unless logistically cumbersome procedures are implemented. These problems have been recognised13, 14 and the use of alternative storage and transport strategies investigated (alcohols or guanidine)15-17. Recently, Kraus et al.18 introduced a method to collect, store and transport AIV samples, based on a special filter paper. FTA cards19 preserve RNA on a dry storage basis20 and render pathogens inactive upon contact21. This study showed that FTA cards can be used to detect AIV RNA in reverse-transcription PCR and that the resulting cDNA could be sequenced and virus genes and determined.
In the study of Kraus et al.18 a laboratory isolate of AIV was used, and samples were handled individually. In the extension presented here, faecal samples from wild birds from the duck trap at the Ottenby Bird Observatory (SE Sweden) were tested directly to illustrate the usefulness of the methods under field conditions. Catching of ducks and sample collection by cloacal swabs is demonstrated. The current protocol includes up-scaling of the work flow from single tube handling to a 96-well design. Although less sensitive than the traditional methods, the method of FTA cards provides an excellent supplement to large surveillance schemes. It allows collection and analysis of samples from anywhere in the world, without the need to maintaining a cool chain or safety regulations with respect to shipping of hazardous reagents, such as alcohol or guanidine.

A method to preserve, detect and sequence RNA from Avian Influenza Viruses was validated and extended using natural faecal samples from birds. This technique removes the necessity of maintaining a cool chain and handling of infectious viruses and can be applied in a 96-well high-throughput setup.

Avian Influenza Viruses (AIVs) infect many mammals, including humans1. These AIVs are diverse in their natural hosts, harboring almost all possible viral ...

Detection of Infectious Virus from Field-collected Mosquitoes by Vero Cell Culture Assay

Mosquitoes transmit a number of distinct viruses including important human pathogens such as West Nile virus, dengue virus, and chickungunya virus. Many of these viruses have intensified in their endemic ranges and expanded to new territories, necessitating effective surveillance and control programs to respond to these threats. One strategy to monitor virus activity involves collecting large numbers of mosquitoes from endemic sites and testing them for viral infection. In this article, we describe how to handle, process, and screen field-collected mosquitoes for infectious virus by Vero cell culture assay. Mosquitoes are sorted by trap location and species, and grouped into pools containing &le;50 individuals. Pooled specimens are homogenized in buffered saline using a mixer-mill and the aqueous phase is inoculated onto confluent Vero cell cultures (Clone E6). Cell cultures are monitored for cytopathic effect from days 3-7 post-inoculation and any viruses grown in cell culture are identified by the appropriate diagnostic assays. By utilizing this approach, we have isolated 9 different viruses from mosquitoes collected in Connecticut, USA, and among these, 5 are known to cause human disease. Three of these viruses (West Nile virus, Potosi virus, and La Crosse virus) represent new records for North America or the New England region since 1999. The ability to detect a wide diversity of viruses is critical to monitoring both established and newly emerging viruses in the mosquito population.

We describe a method to process and screen field-collected mosquitoes for a diversity of viruses by Vero cell culture assay. By employing this technique, we have detected 9 different viruses from 4 taxonomic families in mosquitoes collected in Connecticut.

Mosquitoes transmit a number of distinct viruses including important human pathogens such as West Nile virus, dengue virus, and chickungunya virus. Many ...

Saliva, Salivary Gland, and Hemolymph Collection from Ixodes scapularis Ticks

Ticks are found worldwide and afflict humans with many tick-borne illnesses. Ticks are vectors for pathogens that cause Lyme disease and tick-borne relapsing fever (Borrelia spp.), Rocky Mountain Spotted fever (Rickettsia rickettsii), ehrlichiosis (Ehrlichia chaffeensis and E. equi), anaplasmosis (Anaplasma phagocytophilum), encephalitis (tick-borne encephalitis virus), babesiosis (Babesia spp.), Colorado tick fever (Coltivirus), and tularemia (Francisella tularensis) 1-8. To be properly transmitted into the host these infectious agents differentially regulate gene expression, interact with tick proteins, and migrate through the tick 3,9-13. For example, the Lyme disease agent, Borrelia burgdorferi, adapts through differential gene expression to the feast and famine stages of the tick's enzootic cycle 14,15. Furthermore, as an Ixodes tick consumes a bloodmeal Borrelia replicate and migrate from the midgut into the hemocoel, where they travel to the salivary glands and are transmitted into the host with the expelled saliva 9,16-19.
As a tick feeds the host typically responds with a strong hemostatic and innate immune response 11,13,20-22. Despite these host responses, I. scapularis can feed for several days because tick saliva contains proteins that are immunomodulatory, lytic agents, anticoagulants, and fibrinolysins to aid the tick feeding 3,11,20,21,23. The immunomodulatory activities possessed by tick saliva or salivary gland extract (SGE) facilitate transmission, proliferation, and dissemination of numerous tick-borne pathogens 3,20,24-27. To further understand how tick-borne infectious agents cause disease it is essential to dissect actively feeding ticks and collect tick saliva. This video protocol demonstrates dissection techniques for the collection of hemolymph and the removal of salivary glands from actively feeding I. scapularis nymphs after 48 and 72 hours post mouse placement. We also demonstrate saliva collection from an adult female I. scapularis tick.

Saliva, Salivary Gland, and Hemolymph Collection from Ixodes scapularis Ticks

The collection of infected tick hemolymph, salivary glands, and saliva is important to study how tick-borne pathogens cause disease. In this protocol we demonstrate how to collect hemolymph and salivary glands from feeding Ixodes scapularis nymphs. We also demonstrate saliva collection from female I. scapularis adults.

Ticks are found worldwide and afflict humans with many tick-borne illnesses. Ticks are vectors for pathogens that cause Lyme disease and tick-borne ...

Establishing a Liquid-covered Culture of Polarized Human Airway Epithelial Calu-3 Cells to Study Host Cell Response to Respiratory Pathogens In vitro

The apical and basolateral surfaces of airway epithelial cells demonstrate directional responses to pathogen exposure in vivo. Thus, ideal in vitro models for examining cellular responses to respiratory pathogens polarize, forming apical and basolateral surfaces. One such model is differentiated normal human bronchial epithelial cells (NHBE). However, this system requires lung tissue samples, expertise isolating and culturing epithelial cells from tissue, and time to generate an air-liquid interface culture.
Calu-3 cells, derived from a human bronchial adenocarcinoma, are an alternative model for examining the response of proximal airway epithelial cells to respiratory insult1, pharmacological compounds2-6, and bacterial7-9 and viral pathogens, including influenza virus, rhinovirus and severe acute respiratory syndrome - associated coronavirus10-14. Recently, we demonstrated that Calu-3 cells are susceptible to respiratory syncytial virus (RSV) infection in a manner consistent with NHBE15,16 . Here, we detail the establishment of a polarized, liquid-covered culture (LCC) of Calu-3 cells, focusing on the technical details of growing and culturing Calu-3 cells, maintaining cells that have been cultured into LCC, and we present the method for performing respiratory virus infection of polarized Calu-3 cells.
To consistently obtain polarized Calu-3 LCC, Calu-3 cells must be carefully subcultured before culturing in Transwell inserts. Calu-3 monolayer cultures should remain below 90% confluence, should be subcultured fewer than 10 times from frozen stock, and should regularly be supplied with fresh medium. Once cultured in Transwells, Calu-3 LCC must be handled with care. Irregular media changes and mechanical or physical disruption of the cell layers or plates negatively impact polarization for several hours or days. Polarization is monitored by evaluating trans-epithelial electrical resistance (TEER) and is verified by evaluating the passive equilibration of sodium fluorescein between the apical and basolateral compartments17,18 . Once TEER plateaus at or above 1,000 &Omega;&times;cm2, Calu-3 LCC are ready to use to examine cellular responses to respiratory pathogens.

Establishing a Liquid-covered Culture of Polarized Human Airway Epithelial Calu-3 Cells to Study Host Cell Response to Respiratory Pathogens In vitro

The findings and conclusions in this report are those of the authors and do not necessarily represent the views of the Centers for Disease Control and Prevention.

The  apical and basolateral surfaces of airway epithelial cells demonstrate  directional responses to pathogen exposure in  vivo. Thus, ideal in vitro ...

Using Fluorescent Proteins to Monitor Glycosome Dynamics in the African Trypanosome

Trypanosoma brucei is a kinetoplastid parasite that causes human African trypanosomiasis (HAT), or sleeping sickness, and a wasting disease, nagana, in cattle1. The parasite alternates between the bloodstream of the mammalian host and the tsetse fly vector. The composition of many cellular organelles changes in response to these different extracellular conditions2-5.
Glycosomes are highly specialized peroxisomes in which many of the enzymes involved in glycolysis are compartmentalized. Glycosome composition changes in a developmental and environmentally regulated manner4-11. Currently, the most common techniques used to study glycosome dynamics are electron and fluorescence microscopy; techniques that are expensive, time and labor intensive, and not easily adapted to high throughput analyses.
To overcome these limitations, a fluorescent-glycosome reporter system in which enhanced yellow fluorescent protein (eYFP) is fused to a peroxisome targeting sequence (PTS2), which directs the fusion protein to glycosomes12, has been established. Upon import of the PTS2eYFP fusion protein, glycosomes become fluorescent. Organelle degradation and recycling results in the loss of fluorescence that can be measured by flow cytometry. Large numbers of cells (5,000 cells/sec) can be analyzed in real-time without extensive sample preparation such as fixation and mounting. This method offers a rapid way of detecting changes in organelle composition in response to fluctuating environmental conditions.

Glycosome dynamics in African trypanosomes are difficult to study by traditional cell biology techniques such as electron and fluorescence microscopy. As a means of observing dynamic organelle behavior, a fluorescent-organelle reporter system has been used in conjunction with flow cytometry to monitor real-time glycosome dynamics in live parasites.

Trypanosoma brucei is a kinetoplastid parasite that causes human African trypanosomiasis (HAT), or sleeping sickness, and a wasting disease, nagana, in ...

Intranasal Administration of Recombinant Influenza Vaccines in Chimeric Mouse Models to Study Mucosal Immunity

Vaccines are one of the greatest achievements of mankind, and have saved millions of lives over the last century. Paradoxically, little is known about the physiological mechanisms that mediate immune responses to vaccines perhaps due to the overall success of vaccination, which has reduced interest into the molecular and physiological mechanisms of vaccine immunity. However, several important human pathogens including influenza virus still pose a challenge for vaccination, and may benefit from immune-based strategies. 
Although influenza reverse genetics has been successfully applied to the generation of live-attenuated influenza vaccines (LAIVs), the addition of molecular tools in vaccine preparations such as tracer components to follow up the kinetics of vaccination in vivo, has not been addressed. In addition, the recent generation of mouse models that allow specific depletion of leukocytes during kinetic studies has opened a window of opportunity to understand the basic immune mechanisms underlying vaccine-elicited protection. Here, we describe how the combination of reverse genetics and chimeric mouse models may help to provide new insights into how vaccines work at physiological and molecular levels, using as example a recombinant, cold-adapted, live-attenuated influenza vaccine (LAIV). We utilized laboratory-generated LAIVs harboring cell tracers as well as competitive bone marrow chimeras (BMCs) to determine the early kinetics of vaccine immunity and the main physiological mechanisms responsible for the initiation of vaccine-specific adaptive immunity. In addition, we show how this technique may facilitate gene function studies in single animals during immune responses to vaccines. We propose that this technique can be applied to improve current prophylactic strategies against pathogens for which urgent medical countermeasures are needed, for example influenza, HIV, Plasmodium, and hemorrhagic fever viruses such as Ebola virus.

There is an overall lack of knowledge about how vaccines work. Here we propose the combined use of reverse genetics and bone marrow chimeric mice to gain insight into the early host immune responses to vaccines with a special focus on dendritic cells and T cell immunity.

Vaccines are one of the greatest achievements of mankind, and have saved millions of lives over the last century. Paradoxically, little is known about the ...

Fabricating Optical-quality Glass Surfaces to Study Macrophage Fusion

Visualizing the formation of multinucleated giant cells (MGCs) from living specimens has been challenging due to the fact that most live imaging techniques require propagation of light through glass, but on glass macrophage fusion is a rare event. This protocol presents the fabrication of several optical-quality glass surfaces where adsorption of compounds containing long-chain hydrocarbons transforms glass into a fusogenic surface. First, preparation of clean glass surfaces as starting material for surface modification is described. Second, a method is provided for the adsorption of compounds containing long-chain hydrocarbons to convert non-fusogenic glass into a fusogenic substrate. Third, this protocol describes fabrication of surface micropatterns that promote a high degree of spatiotemporal control over MGC formation. Finally, fabricating glass bottom dishes is described. Examples of use of this in vitro cell system as a model to study macrophage fusion and MGC formation are shown.

This protocol describes the fabrication of optical-quality glass surfaces adsorbed with compounds containing long-chain hydrocarbons that can be used to monitor macrophage fusion of living specimens and enables super-resolution microscopy of fixed specimens.

Visualizing the formation of multinucleated giant cells (MGCs) from living specimens has been challenging due to the fact that most live imaging ...

Immunohistochemistry Test for the Lyssavirus Antigen Detection from Formalin-Fixed Tissues

One of the primary diagnostic modalities for rabies is the detection of viral ribonucleoprotein (RNP) complex (antigen) in the infected tissue samples. While the direct fluorescent antibody (DFA) test or the direct rapid immunohistochemical test (DRIT) are most commonly utilized for the antigen detection, both tests require fresh and/or frozen tissues for impressions on slides prior to the antigen detection using antibodies. If samples are collected and fixed in formalin, neither test is optimal for the antigen detection, however, testing can be performed by conventional immunohistochemistry (IHC) after embedding in paraffin blocks and sectioning. With this IHC method, tissues are stained with anti-rabies antibodies, sections are deparaffinized, antigen retrieved by partial proteolysis or other methods, and incubated with primary and secondary antibodies. Antigens are stained using horseradish peroxidase / amino ethyl carbazole and counterstained with hematoxylin for the visualization using a light microscope. In addition to the specific antigen detection, formalin fixation offers other advantages like the determination of histological changes, relaxed conditions for specimen storage and transport (under ambient temperatures), ability to test retrospective cases and improved biological safety through the inactivation of infectious agents.

Here, we present an immunohistochemistry test protocol for the detection of rabies virus antigen as an alternative diagnostic test for formalin-fixed tissues.

One of the primary diagnostic modalities for rabies is the detection of viral ribonucleoprotein (RNP) complex (antigen) in the infected tissue samples. ...