We wish to thank M. Scullion and R. Crisp of Idaho Technologies for technical support and the USGS Western Ecological Research Center for 
funding (S. Schwarzbach) and assistance (K. Spragens, T. Graham). This research was performed under the auspices of the Center for Innovative Technology - 
Institute for Defense and Homeland Security (www.idhs.org), in support of the Department of Defense and Air Force Research Laboratory. Animal handling 
followed protocols approved by the Animal Care and Use Committee of the U.S. Geological Survey Western Ecological Research Center and permits of the U.S. Geological Survey
Bird Banding Laboratory. Any use of trade, product, or firm names in this publication is for descriptive purposes only and does not imply endorsement by the 
U. S. government.

1. Wild bird capture using mist nets
<ol>
<li>For shorebird capture, set up mist nets at an active foraging site such as a marsh, shoreline, or mud flat.</li>
<li>Slide trammel line loops of one end of the mist net around the pole and insert pole vertically into mud.</li>
<li>Stretch out net, insert second pole through trammel loops at other end of mist net and vertically insert pole into mud, making sure that the trammel lines are taught.</li>
<li>Once a bird is captured, extract the bird from the net and return to the banding station.</li>
</ol>
2. Cloacal swab sampling
<ol>
<li>Collect cloacal swabs immediately after capture</li>
<li>Locate the cloaca and use fingertips to push back surrounding feathers</li>
<li>Unwrap a sterile Dacron- or Polyester-tipped swab, stem end first, and moisten the tip with sterile viral transport media for greater ease in swabbing. After moistening, the entire head of the swab is gently inserted into the cloaca. Swab the inside circumference of the cloaca by slowly twirling the swab and remove the swab from cloaca.</li>
<li>Inserting the swab tip directly into the viral transport media sample vial; rotate the swab shaft between the thumb and forefinger while the swab is in the media. Have an assistant complete the procedure by pulling the tip of the swab back from the bottom of the vial ~ 1 cm and cut the shaft of the swab with a pair of scissors so the swab tip remains in the vial and the shaft does not interfere with cap closure. Cap vial tightly. Disinfect the scissors with alcohol in between cutting swab shafts.</li>
<li>Place the vials in a freezer box containing either dry ice or ice packs.</li>
</ol>
3. Oropharyngeal swab sampling
<ol start="1">
<li>Gently open the bird's beak so that the choanal slit is visible</li>
<li>Unwrap a clean swab from the package, stem end first, and moisten the tip with viral transport media for greater ease in swabbing</li>
<li>Without touching the tip to any other surfaces, insert it into the mouth and swab the choanal opening on the roof of the mouth and continue back towards the oropharyngeal or throat region</li>
<li>Place the swab tip directly into the viral transport media vial. Rotate the swab shaft between the thumb and forefinger while the swab is in the media. As above, have an assistant pull the tip of the swab back from the bottom of the vial ~ 1 cm and cut the shaft of the swab with a pair of scissors so the swab tip remains in the vial and the shaft does not interfere with cap closure. Cap vial tightly. Disinfect the scissors with alcohol in between cutting swab shafts.</li>
<li>Place the vials in a freezer box containing dry ice or ice packs.</li>
</ol>
4. 	Bird handling and release
<ol start="1">
	<li>Return bird to the site of capture and release in a timely manner (&lt;20mins). Handlers should be aware of the principles of animal welfare and be alert for signs of bird suffering during sampling (i.e. abrasions, capture myopathy and hypo- or hyperthermia). A basic first aid kit should be included in the equipment list.</li>
</ol>
5. Testing facilities
<ol start="1">
<li>Samples can be tested in any enclosed space such as a research trailer or tent, with a power supply (voltage-regulated power generator) to run the PCR unit and attached computer.</li>
</ol>
6. RNA extraction
<ol start="1">
<li>RNA extraction was performed with the RNeasy Mini kit with minor modifications to the manufacturer's instructions following Spackman et al. 1</li>
<li>Vortex the swab medium for 3-5 s and transfer 350 &mu;l to a 2 ml microcentrifuge tube. Note: Keep remainder sample on ice in case PCR-positive samples need to be rescreened or H5 test needs to be run.</li>
<li>Add 350 &mu;l of RLT buffer (with &beta;-me) and vortex for 5 s.</li>
<li>Add 350 &mu;l of RNA grade 70% ethanol.</li>
<li>Centrifuge at 5,000 x g for 5 min.</li>
<li>Add 600 &mu;l of the supernatant to a spin column placed in a 2 ml collection tube. Retain the remaining sample in test tube rack.</li>
<li>Centrifuge for 20 s at 10,000 x g and discard flow-through.</li>
<li>Repeat steps 17-18 until all the sample has been passed through spin column.</li>
<li>Place spin column in a clean 2 ml collection tube.</li>
<li>Add 700 &mu;l of RW1 buffer to the spin column and centrifuge 25 s at 10,000 x g. Discard the flow-through.</li>
<li>Add 500 &mu;l RPE buffer to the spin column and centrifuge for 25 s at 10,000 x g. Discard the flow-through.</li>
<li>Repeat step 22 for a total of two washes with RPE buffer.</li>
<li>Centrifuge the spin column for an additional 2 minutes at 14,000 x g. Discard collection tube.</li>
<li>Place the spin column in a clean 1.5 ml microcentrifuge tube and add 50&mu;l of nuclease-free water. Incubate at room temperature for 60 s.</li>
<li>Elute RNA by spinning for 60 s at 10,000 x g for 60 s. Place purified sample on ice.</li>
</ol>
7. Preparing negative controls
<ol start="1">
<li>Lyophilized reagents were prepared for use in the PCR assay according to instructions from the Idaho Technologies Freeze-Dried Reagent Detection Kit for TaqMan Probes Influenza A Target 1 (ASAY-ASY-0109).</li>
<li>Centrifuge negative control reagent vial (red) for 3 s to ensure pellets are at the bottom.</li>
<li>Remove cap, visually make sure pellets are at the bottom.</li>
<li>Add 20 &mu;l of 2 X Reconstitution Buffer and 20 &mu;l Reagent Grade Water.</li>
<li>Recap and vortex for 5 s at maximum speed.</li>
<li>Centrifuge for 3 s to bring liquid to bottom of vial.</li>
<li>Visually check the pellet has rehydrated, if not repeat steps 31-32.</li>
<li>Pipette 19 &mu;l of the hydrated mixture into a Lightcycler capillary tube, repeat pipette step into a second capillary to obtain a duplicate.</li>
</ol>
8. Preparing test samples
<ol start="1">
<li>Centrifuge the unknown reagent vial (blue) for 3 sec to ensure pellets are at the bottom.</li>
<li>Remove cap, visually make sure pellets are at the bottom.</li>
<li>Add 40 &mu;l of purified RNA.</li>
<li>Recap and vortex for 5 s at maximum speed.</li>
<li>Centrifuge for 3 s to bring liquid to bottom of vial.</li>
<li>Visually check the pellet has rehydrated, if not repeat steps 38-39. </li>
<li>Pipette 19 &mu;l of hydrated mixture into a Lightcycler capillary tube, repeat pipette step into a second capillary to obtain a duplicate.</li>
</ol>
9. Preparing positive controls
<ol start="1">
<li>Centrifuge the positive control reagent vial (green) for 3 s to ensure pellets are at the bottom.</li>
<li>Remove cap, visually make sure pellets are at the bottom.</li>
<li>Add 20 &mu;l of 2 X Reconstitution Buffer and 20 &mu;l Reagent Grade Water.</li>
<li>Recap and vortex for 5 s at maximum speed.</li>
<li>Centrifuge for 3 s to bring liquid to bottom of vial.</li>
<li>Visually check the pellet has rehydrated, if not repeat steps 45-46.</li>
<li>Pipette 19 &mu;l of the hydrated mixture into a Lightcycler capillary tube, repeat pipette step into a second capillary to obtain a duplicate.</li>
</ol>
10. Centrifuging the capillary tubes
<ol start="1">
<li>Once all of the capillary tubes for the batch have been prepared load them into the mini-centrifuge fitted with capillary adapters.</li>
<li>Centrifuge at the lowest setting by holding the pulse button down for 1 s. This transfers liquid to the bottom in preparation for testing.</li>
</ol>
11. Operating the RAPID
<ol start="1">
<li>Create a New file. Label positive control, negative control, and samples (by their ID number).</li>
<li>Program 1: RNA Sample Preparation Hold
<ol>
<li>Cycles = 1</li>
<li>Analysis Mode = None</li>
<li>Target Temperature = 40</li>
<li>Incubation Time = 30 min</li>
<li>Temperature Transition Rate = 20&deg;C/s (default)</li>
<li>Secondary Temperature Target = 0 (default)</li>
<li>Step Size = 0 (default)</li>
<li>Step Delay = 0 (default)</li>
<li>Acquisition Mode = None (default)</li>
</ol>
</li>
<li>Program 2: RNA Denaturation
<ol>
<li>Cycles = 1</li>
<li>Analysis Mode = None</li>
<li>Target Temperature = 94</li>
<li>Incubation Time = 120s</li>
<li>Temperature Transition Rate = 20&deg;C/s (default)</li>
<li>Secondary Temperature Target = 0 (default)</li>
<li>Step Size = 0 (default)</li>
<li>Step Delay = 0 (default)</li>
<li>Acquisition Mode = None (default)</li>
</ol>
</li>
<li>Program 3: RNA Amplification
<ol>
<li>Cycles = 45</li>
<li>Analysis Mode = Auto Analysis</li>
<li>Target Temperature, Segment 1 = 94; Segment 2 = 60</li>
<li>Incubation Time, Segment 1 = 0 s, Segment 2 = 20 s</li>
<li>Temperature Transition Rate, Seg 1 &amp; 2 = 20&deg;C/s (default)</li>
<li>Secondary Temperature Target, Seg 1 &amp; 2 = 0 (default)</li>
<li>Step Size, Seg 1 &amp; 2 = 0 (default)</li>
<li>Step Delay, Seg 1 &amp; 2 = 0 (default)</li>
<li>Acquisition Mode, Segment 1 = None; Segment 2 = single</li>
</ol>
</li>
<li>Fluorescence Parameters
<ol>
<li>Display Mode: fluorescence channel 2 (Ch2/1) 
Gains:
<ol>
<li>Ch 1 (F1) = 1</li>
<li>Ch 2 (F2) = 8</li>
<li>Ch 3 (F3) = 1</li>
</ol>
</li>
<li>To display the real-time fluorescence data for each sample, select the sample of interest and click the 'Show Graph' button at the bottom of the screen</li>
</ol>
</li>
<li>For each capillary, the results include the following:
<ul>
<li># -- identifies the carousel position of the carousel</li>
<li>Sample name - provides the name of the sample</li>
<li>Control - indentifies the sample type for each sample</li>
<li>Score - calculated using the relationship between the sample's fluorescence and the level of background fluorescence in the sample</li>
<li>Result - displays the overall result for the sample</li>
</ul>
</li>
</ol>
12. Representative results:
Possible results include the following: 
Present: A red 'present' call for a test/unknown sample indicates that the target was identified in that sample and the controls were successful. 
Not detected: A green 'not detected' call for a test/unknown sample indicates that the target was not identified in that sample and the test controls were successful. 
Please repeat: A 'please repeat' indicates that the positive or negative control failed.
An example of a successful analysis by the RAPID 7200 is shown in Figure 4. The positive control is amplified and generates detectable fluorescence (y-axis) at approximately 25 cycles (x-axis). A cycle threshold (CT) > 35 is the agreed cut-off for most AIV reference laboratories. Therefore, the positive controls for this method produced a fluorescent signal within the accepted number of cycles (0-35) for AIV detection. In contrast, the negative control does not generate a fluorescent signal even after 45 cycles. Similarly, the twelve samples collected from western sandpipers did not generate a fluorescent signal indicating that the birds were negative for AIV. Follow-up testing of all positive samples in a traditional laboratory is encouraged to ensure that no false positives are erroneously produced.
<img src="/files/ftp_upload/2829/2829fig1.jpg" alt="Figure 1" /> 
Figure 1. Shorebird capture using mist nets.
<img src="/files/ftp_upload/2829/2829fig2.jpg" alt="Figure 2" /> 
Figure 2. Collection of cloacal and oropharyngeal samples from least sandpipers.
<img src="/files/ftp_upload/2829/2829fig3.jpg" alt="Figure 3" /> 
Figure 3. Programming the RAPID 7200 for RNA amplification.
<img src="/files/ftp_upload/2829/2829fig4.jpg" alt="Figure 4" /> 
Figure 4. Fluorogram generated by the RAPID 7200 portable rRT-PCR showing how positive and negative controls should appear in a successful assay. <a href="https://www.jove.com/files/ftp_upload/2829/2829fig4large.jpg">Click here to view the full sized image</a>

<ol>
	<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+transcriptase+PCR+assay+for+type+A+influenza+virus+and+the+avian+H5+and+H7+hemagglutinin+subtypes.">Development of a real-time reverse transcriptase PCR assay for type A influenza virus and the avian H5 and H7 hemagglutinin subtypes.</a> J Clin Microbiol. 40, 3256-3260 (2002).</li><li>Takekawa, J. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Field+detection+of+avian+influenza+virus+in+wild+birds:+evaluation+of+a+portable+rRT-PCR+system+and+freeze-dried+reagents.">Field detection of avian influenza virus in wild birds: evaluation of a portable rRT-PCR system and freeze-dried reagents.</a> J Virol Methods. 166, 92-97 (2010).</li><li>Das, A., Spackman, E., Senne, D., Pedersen, J., Suarez, D. 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No conflicts of interest declared.

The method of rapid diagnosis presented here facilitates time-efficient and accurate testing of wild bird samples for surveillance of AIV. 
The much less stringent specimen storage requirements of portable rRT-PCR are suitable for remote situations where maintenance of a cold chain may be impractical if liquid nitrogen shippers or dry ice is not available. In addition, we found that sample analysis with freeze-dried reagents was straightforward enough to be performed by field biologists with minimal knowledge of laborato...

<table border="1">
<tbody>
 <tr>
 <th>Name of the reagent</th>
 <th>Company</th>
 <th>Catalogue Number</th>
 <th>Comments (optional)</th>
 </tr>
 <tr>
 <td>RNeasy mini spin column</td>
 <td>Qiagen</td>
 <td>74106</td>
 <td>Included in RNeasy Mini Kit</td>
 </tr>
 <tr>
 <td>Collection tubes (1.5 &amp; 2 mL)</td>
 <td>Qiagen</td>
 <td>74106</td>
 <td>Included in RNeasy Mini Kit</td>
 </tr>
 <tr>
 <td>Buffer RLT</td>
 <td>Qiagen</td>
 <td>74106</td>
 <td>Included in RNeasy Mini Kit</td>
 </tr>
 <tr>
 <td>Buffer RW1</td>
 <td>Qiagen</td>
 <td>74106</td>
 <td>Included in RNeasy Mini Kit</td>
 </tr>
 <tr>
 <td>Buffer RPE</td>
 <td>Qiagen</td>
 <td>74106</td>
 <td>Included in RNeasy Mini Kit</td>
 </tr>
 <tr>
 <td>RNase-free water</td>
 <td>Qiagen</td>
 <td>74106</td>
 <td>Included in RNeasy Mini Kit</td>
 </tr>
 <tr>
 <td>14.3 M &beta;-mercaptoethanol solution</td>
 <td>Fisher Scientific</td>
 <td>BP176100</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>100% ethanol</td>
 <td>Fisher Scientific</td>
 <td>NC9602322</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Vortex Genie 2, 120V</td>
 <td>Scientific Industries</td>
 <td>SI-0236</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Taqman Influenza A Target 1 (Hydrolysis Probe)</td>
 <td>Idaho Technologies</td>
 <td>ASAY-ASY-0109</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Lightcycler 20ml capillary tubes</td>
 <td>Roche Applied Science</td>
 <td>04929292001</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Micro-centrifuge with rotator for 2 ml tubes </td>
 <td>Idaho Technologies</td>
 <td>&nbsp;</td>
 <td>Included in RAPID kit</td>
 </tr>
 <tr>
 <td>Ruggedized Advanced Pathogen Identification Device (RAPID) 7200</td>
 <td>Idaho Technologies</td>
 <td>&nbsp;</td>
 <td>Included in RAPID kit</td>
 </tr>
 <tr>
 <td>Pentium-based laptop with Windows XP Professional</td>
 <td>Idaho Technologies</td>
 <td>&nbsp;</td>
 <td>Included in RAPID kit</td>
 </tr>
 <tr>
 <td>Lightcycler Data Analysis software</td>
 <td>Idaho Technologies</td>
 <td>&nbsp;</td>
 <td>Included in RAPID kit</td>
 </tr>
 </tbody>
</table>

rapid diagnosis of avian influenza virus in wild birds: use of a portable rrt-pcr and freeze-dried reagents in the field

Wild birds have been implicated in the spread of highly pathogenic avian influenza (HPAI) of the H5N1 subtype, prompting surveillance along migratory flyways. Sampling of wild birds for avian influenza virus (AIV) is often conducted in remote regions, but results are often delayed because of the need to transport samples to a laboratory equipped for molecular testing. Real-time reverse transcriptase polymerase chain reaction (rRT-PCR) is a molecular technique that offers one of the most accurate and sensitive methods for diagnosis of AIV. The previously strict lab protocols needed for rRT-PCR are now being adapted for the field. Development of freeze-dried (lyophilized) reagents that do not require cold chain, with sensitivity at the level of wet reagents has brought on-site remote testing to a practical goal.
Here we present a method for the rapid diagnosis of AIV in wild birds using an rRT-PCR unit (Ruggedized Advanced Pathogen Identification Device or RAPID, Idaho Technologies, Salt Lake City, UT) that employs lyophilized reagents (Influenza A Target 1 Taqman; ASAY-ASY-0109, Idaho Technologies). The reagents contain all of the necessary components for testing at appropriate concentrations in a single tube: primers, probes, enzymes, buffers and internal positive controls, eliminating errors associated with improper storage or handling of wet reagents. The portable unit performs a screen for Influenza A by targeting the matrix gene and yields results in 2-3 hours. Genetic subtyping is also possible with H5 and H7 primer sets that target the hemagglutinin gene.
The system is suitable for use on cloacal and oropharyngeal samples collected from wild birds, as demonstrated here on the migratory shorebird species, the western sandpiper (Calidrus mauri) captured in Northern California. Animal handling followed protocols approved by the Animal Care and Use Committee of the U.S. Geological Survey Western Ecological Research Center and permits of the U.S. Geological Survey Bird Banding Laboratory. The primary advantage of this technique is to expedite diagnosis of wild birds, increasing the chances of containing an outbreak in a remote location. On-site diagnosis would also prove useful for identifying and studying infected individuals in wild populations. The opportunity to collect information on host biology (immunological and physiological response to infection) and spatial ecology (migratory performance of infected birds) will provide insights into the extent to which wild birds can act as vectors for AIV over long distances.

Watch this Scientific Journal Video about Rapid Diagnosis of Avian Influenza Virus in Wild Birds: Use of a Portable rRT-PCR and Freeze-dried Reagents in the Field at JoVE.com

Rapid Diagnosis of Avian Influenza Virus in Wild Birds: Use of a Portable rRT-PCR and Freeze-dried Reagents in the Field

This study describes diagnosis of avian influenza in wild birds using a portable rRT-PCR system. The method takes advantage of freeze-dried reagents to screen wild birds in a non-laboratory setting, typical of an outbreak scenario. Use of molecular tools provides accurate and sensitive alternatives for rapid diagnosis.

Wild birds have been implicated in the spread of highly pathogenic avian influenza (HPAI) of the H5N1 subtype, prompting surveillance along migratory ...

rapid-diagnosis-avian-influenza-virus-wild-birds-use-portable-rrt-pcr

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JoVE Journal

Immunology and Infection

15.6K Views.  USGS Western Ecological Research Center. This study describes diagnosis of avian influenza in wild birds using a portable rRT-PCR system. The method takes advantage of freeze-dried reagents to screen wild birds in a non-laboratory setting, typical of an outbreak scenario. Use of molecular tools provides accurate and sensitive alternatives for rapid diagnosis.

Video: Rapid Diagnosis of Avian Influenza Virus in Wild Birds: Use of a Portable rRT-PCR and Freeze-dried Reagents in the Field

Using Bioluminescent Imaging to Investigate Synergism Between Streptococcus pneumoniae and Influenza A Virus in Infant Mice

During the 1918 influenza virus pandemic, which killed approximately 50 million people worldwide, the majority of fatalities were not the result of infection with influenza virus alone. Instead, most individuals are thought to have succumbed to a secondary bacterial infection, predominately caused by the bacterium Streptococcus pneumoniae (the pneumococcus). The synergistic relationship between infections caused by influenza virus and the pneumococcus has subsequently been observed during the 1957 Asian influenza virus pandemic, as well as during seasonal outbreaks of the virus (reviewed in 1, 2). Here, we describe a protocol used to investigate the mechanism(s) that may be involved in increased morbidity as a result of concurrent influenza A virus and S. pneumoniae infection. We have developed an infant murine model to reliably and reproducibly demonstrate the effects of influenza virus infection of mice colonised with S. pneumoniae. Using this protocol, we have provided the first insight into the kinetics of pneumococcal transmission between co-housed, neonatal mice using in vivo imaging 3.

Using Bioluminescent Imaging to Investigate Synergism Between Streptococcus pneumoniae and Influenza A Virus in Infant Mice

A concurrent infection with influenza A virus is one of the factors implicated in the induction of invasive pneumococcal disease during asymptomatic Streptococcus pneumoniae carriage. Here we describe a mixed infection method using infant mice to investigate the synergism between these two respiratory pathogens.

During the 1918 influenza virus pandemic, which killed approximately 50 million people worldwide, the majority of fatalities were not the result of ...

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 ...

Microwave Assisted Rapid Diagnosis of Plant Virus Diseases by Transmission Electron Microscopy

Investigations of ultrastructural changes induced by viruses are often necessary to clearly identify viral diseases in plants. With conventional sample preparation for transmission electron microscopy (TEM) such investigations can take several days1,2 and are therefore not suited for a rapid diagnosis of plant virus diseases. Microwave fixation can be used to drastically reduce sample preparation time for TEM investigations with similar ultrastructural results as observed after conventionally sample preparation3-5. Many different custom made microwave devices are currently available which can be used for the successful fixation and embedding of biological samples for TEM investigations5-8. In this study we demonstrate on Tobacco Mosaic Virus (TMV) infected Nicotiana tabacum plants that it is possible to diagnose ultrastructural alterations in leaves in about half a day by using microwave assisted sample preparation for TEM. We have chosen to perform this study with a commercially available microwave device as it performs sample preparation almost fully automatically5 in contrast to the other available devices where many steps still have to be performed manually6-8 and are therefore more time and labor consuming. As sample preparation is performed fully automatically negative staining of viral particles in the sap of the remaining TMV-infected leaves and the following examination of ultrastructure and size can be performed during fixation and embedding.

This study describes a method that allows the rapid and clear diagnosis of plant virus diseases in about half a day by using a combination of microwave assisted plant sample preparation for transmission electron microscopy and negative staining methods.

Investigations of ultrastructural changes induced by viruses are often necessary to clearly identify viral diseases in plants. With conventional sample ...

Influenza Virus Propagation in Embryonated Chicken Eggs

Influenza infection is associated with about 36,000 deaths and more than 200,000 hospitalizations every year in the United States. The continuous emergence of new influenza virus strains due to mutation and re-assortment complicates the control of the virus and necessitates the permanent development of novel drugs and vaccines. The laboratory-based study of influenza requires a reliable and cost-effective method for the propagation of the virus. Here, a comprehensive protocol is provided for influenza A virus propagation in fertile chicken eggs, which consistently yields high titer viral stocks. In brief, serum pathogen-free (SPF) fertilized chicken eggs are incubated at 37 &#176;C and 55-60% humidity for 10 &#8211; 11 days. Over this period, embryo development can be easily monitored using an egg candler. Virus inoculation is carried out by injection of virus stock into the allantoic cavity using a needle. After 2 days of incubation at 37 &#176;C, the eggs are chilled for at least 4 hr at 4 &#176;C. The eggshell above the air sac and the chorioallantoic membrane are then carefully opened, and the allantoic fluid containing the virus is harvested. The fluid is cleared from debris by centrifugation, aliquoted and transferred to -80 &#176;C for long-term storage. The large amount (5-10 ml of virus-containing fluid per egg) and high virus titer which is usually achieved with this protocol has made the usage of eggs for virus preparation our favorable method, in particular for in vitro studies which require large quantities of virus in which high dosages of the same virus stock are needed.

Fertile chicken eggs are widely used to produce large amounts of human influenza A virus as they provide a convenient and cost-effective system to prove high yields of virus.

Influenza infection is associated with about 36,000 deaths and more than 200,000 hospitalizations every year in the United States. The continuous ...

Co-immunoprecipitation of the Mouse Mx1 Protein with the Influenza A Virus Nucleoprotein

Studying the interaction between proteins is key in understanding their function(s). A very powerful method that is frequently used to study interactions of proteins with other macromolecules in a complex sample is called co-immunoprecipitation. The described co-immunoprecipitation protocol allows to demonstrate and further investigate the interaction between the antiviral myxovirus resistance protein 1 (Mx1) and one of its viral targets, the influenza A virus nucleoprotein (NP). The protocol starts with transfected mammalian cells, but it is also possible to use influenza A virus infected cells as starting material. After cell lysis, the viral NP protein is pulled-down with a specific antibody and the resulting immune-complexes are precipitated with protein G beads. The successful pull-down of NP and the co-immunoprecipitation of the antiviral Mx1 protein are subsequently revealed by western blotting. A prerequisite for successful co-immunoprecipitation of Mx1 with NP is the presence of N-ethylmaleimide (NEM) in the cell lysis buffer. NEM alkylates free thiol groups. Presumably this reaction stabilizes the weak and/or transient NP&#8211;Mx1 interaction by preserving a specific conformation of Mx1, its viral target or an unknown third component. An important limitation of co-immunoprecipitation experiments is the inadvertent pull-down of contaminating proteins, caused by nonspecific binding of proteins to the protein G beads or antibodies. Therefore, it is very important to include control settings to exclude false positive results. The described co-immunoprecipitation protocol can be used to study the interaction of Mx proteins from different vertebrate species with viral proteins, any pair of proteins, or of a protein with other macromolecules. The beneficial role of NEM to stabilize weak and/or transient interactions needs to be tested for each interaction pair individually.

This co-immunoprecipitation protocol allows to study the interaction between the influenza A virus nucleoprotein and the antiviral Mx1 protein in human cells. The protocol emphasizes the importance of N-ethylmaleimide for successful co-immunoprecipitation of Mx1 and influenza A virus nucleoprotein.

Studying the interaction between proteins is key in understanding their function(s). A very powerful method that is frequently used to study interactions ...

Nasal Wipes for Influenza A Virus Detection and Isolation from Swine

Surveillance for influenza A viruses in swine is critical to human and animal health because influenza A virus rapidly evolves in swine populations and new strains are continually emerging. Swine are able to be infected by diverse lineages of influenza A virus making them important hosts for the emergence and maintenance of novel influenza A virus strains. Sampling pigs in diverse settings such as commercial swine farms, agricultural fairs, and live animal markets is important to provide a comprehensive view of currently circulating IAV strains. The current gold-standard ante-mortem sampling technique (i.e. collection of nasal swabs) is labor intensive because it requires physical restraint of the pigs. Nasal wipes involve rubbing a piece of fabric across the snout of the pig with minimal to no restraint of the animal. The nasal wipe procedure is simple to perform and does not require personnel with professional veterinary or animal handling training. While slightly less sensitive than nasal swabs, virus detection and isolation rates are adequate to make nasal wipes a viable alternative for sampling individual pigs when low stress sampling methods are required. The proceeding protocol outlines the steps needed to collect a viable nasal wipe from an individual pig.

The authors present a protocol to collect swine nasal wipes to detect and isolate influenza A viruses.

Surveillance for influenza A viruses in swine is critical to human and animal health because influenza A virus rapidly evolves in swine populations and ...

Neutrophil Isolation and Analysis to Determine their Role in Lymphoma Cell Sensitivity to Therapeutic Agents

Neutrophils are the most abundant (40% to 75%) type of white blood cells and among the first inflammatory cells to migrate towards the site of inflammation. They are key players in the innate immune system and play major roles in cancer biology. Neutrophils have been proposed as key mediators of malignant transformation, tumor progression, angiogenesis and in the modulation of the antitumor immunity; through their release of soluble factors or their interaction with tumor cells. To characterize the specific functions of neutrophils, a fast and reliable method is coveted for in vitro isolation of neutrophils from human blood. Here, a density gradient separation method is demonstrated to isolate neutrophils as well as mononuclear cells from the blood. The procedure consists of layering the density gradient solution such as Ficoll carefully above the diluted blood obtained from patients diagnosed with chronic lymphocytic leukemia (CLL), followed by centrifugation, isolation of mononuclear layer, separation of neutrophils from RBCsby dextran then lysis of residual erythrocytes. This method has been shown to isolate neutrophils &#8805; 90 % pure. To mimic the tumor microenvironment, 3-dimensional (3D) experiments were performed using basement membrane matrix such as Matrigel. Given the short half-life of neutrophils in vitro, 3D experiments with fresh human neutrophils cannot be performed. For this reason promyelocytic HL60 cells are differentiated along the granulocytic pathway using the differentiation inducers dimethyl sulfoxide (DMSO) and retinoic acid (RA). The aim of our experiments is to study the role of neutrophils on the sensitivity of lymphoma cells to anti-lymphoma agents. However these methods can be generalized to study the interactions of neutrophils or neutrophil-like cells with a large range of cell types in different situations.

Neutrophils are the most abundant type of white blood cells. The isolation of neutrophils from human blood by density gradient separation method and the differentiation of human promyelocytic (HL60) cells along the granulocytic pathway are described here; to test their role on sensitivity of lymphoma cells to anti-lymphoma agents.

Neutrophils are the most abundant (40% to 75%) type of white blood cells and among the first inflammatory cells to migrate towards the site of ...

In Vitro Disassembly of Influenza A Virus Capsids by Gradient Centrifugation

Acid-triggered molecular processes closely control cell entry of many viruses that enter through the endocytic system. In the case of influenza A virus (IAV), virus fusion with the endosomal membrane as well as the subsequent disassembly of the viral capsid, called uncoating, is governed by the ionic conditions inside endocytic vesicles. The early steps in the virus life cycle are hard to study because endosomes cannot be directly accessed experimentally, creating the need for an in vitro approach. Here, we describe a method based on velocity gradient centrifugation of purified virions through a two-layer glycerol gradient, which enables analysis of the IAV core and its stability. The gradient contains a non-ionic detergent (NP-40) in its lower layer to remove the viral membrane by solubilization as the virus sediments toward the bottom. At neutral pH, viral cores are pelleted as stable structures. The major core components, matrix protein (M1) and the viral ribonucleoproteins (vRNPs), can be clearly identified in the pellet fraction by SDS-PAGE. Decreasing the pH to 6.0 or lower in the bottom layer selectively removes M1 from the pellet followed by release of vRNPs at more acidic conditions. Viral protein bands on Coomassie-stained gels can be subjected to densitometric quantification to monitor intermediate states of IAV disassembly. Besides pH, other factors that influence viral core stability can be assessed, such as salt concentration and putative viral uncoating factors, simply by modifying the detergent-containing glycerol layer accordingly. Taken together, the presented technique allows highly reproducible and quantitative analysis of viral uncoating in vitro. It can be applied to other enveloped viruses that undergo complex uncoating processes.

In Vitro Disassembly of Influenza A Virus Capsids by Gradient Centrifugation

Disassembly of influenza A virus cores during virus entry into host cells is a multistep process. We describe an in vitro method to analyze the early stages of viral uncoating. In this approach, velocity gradient centrifugation is used to biochemically dissect the steps that initiate uncoating under defined conditions.

Acid-triggered molecular processes closely control cell entry of many viruses that enter through the endocytic system. In the case of influenza A virus ...

Evaluation of the Efficacy And Toxicity of RNAs Targeting HIV-1 Production for Use in Gene or Drug Therapy

Small RNA therapies targeting post-integration steps in the HIV-1 replication cycle are among the top candidates for gene therapy and have the potential to be used as drug therapies for HIV-1 infection. Post-integration inhibitors include ribozymes, short hairpin (sh) RNAs, small interfering (si) RNAs, U1 interference (U1i) RNAs and RNA aptamers. Many of these have been identified using transient co-transfection assays with an HIV-1 expression plasmid and some have advanced to clinical trials. In addition to measures of efficacy, small RNAs have been evaluated for their potential to affect the expression of human RNAs, alter cell growth and/or differentiation, and elicit innate immune responses. In the protocols described here, a set of transient transfection assays designed to evaluate the efficacy and toxicity of RNA molecules targeting post-integration steps in the HIV-1 replication cycle are described. We have used these assays to identify new ribozymes and optimize the format of shRNAs and siRNAs targeting HIV-1 RNA. The methods provide a quick set of assays that are useful for screening new anti-HIV-1 RNAs and could be adapted to screen other post-integration inhibitors of HIV-1 replication.

Methods to evaluate the efficacy and toxicity of RNA molecules targeting post-integration steps of the HIV-1 replication cycle are described. These methods are useful for screening new molecules and optimizing the format of existing ones.

Small RNA therapies targeting post-integration steps in the HIV-1 replication cycle are among the top candidates for gene therapy and have the potential ...

Influenza A Virus Studies in a Mouse Model of Infection

Influenza viruses cause over 500,000 deaths worldwide1 and are associated with an annual cost of 12 - 14 billion USD in the United States alone considering direct medical and hospitalization expenses and work absenteeism2. Animal models are crucial in Influenza A virus (IAV) studies to evaluate viral pathogenesis, host-pathogen interactions, immune responses, and the efficacy of current and/or novel vaccine approaches as well as antivirals. Mice are an advantageous small animal model because their immune system is evolutionarily similar to that found in humans, they are available from commercial vendors as genetically identical subjects, there are multiple strains that can be exploited to evaluate the genetic basis of infections, and they are relatively inexpensive and easy to manipulate. To recapitulate IAV infection in humans via the airways, mice are first anesthetized prior to intranasal inoculation with infectious IAVs under proper biosafety containment. After infection, the pathogenesis of IAVs is determined by monitoring daily the morbidity (body weight loss) and mortality (survival) rate. In addition, viral pathogenesis can also be evaluated by assessing virus replication in the upper (nasal mucosa) or lower (lungs) respiratory tract of infected mice. Humoral responses upon IAV infection can be rapidly evaluated by non-invasive bleeding and secondary antibody detection assays aimed at detecting the presence of total or neutralizing antibodies. Here, we describe the common methods used to infect mice intranasally (i.n) with IAV and evaluate pathogenesis, humoral immune responses and protection efficacy.

Influenza A viruses (IAVs) are important human respiratory pathogens. To understand the pathogenicity of IAVs and to perform preclinical testing of novel vaccine approaches, animal models mimicking human physiology are required. Here, we describe techniques to evaluate IAV pathogenesis, humoral responses and vaccine efficacy using a mouse model of infection.