1. Whitefly Colony Maintenance
<ol>
	<li>Environmental Conditions: Whitefly colonies should be maintained in a controlled growth room. Control of relative humidity, temperature, photoperiod, and light intensity are essential for optimal colony growth (Figure 1). A temperature of 28 &#176;C, 30-50% relative humidity, and a 14 hr photoperiod will yield a colony that develops from egg to adult emergence in 18 days (this time varies with the plant host). Relative humidity should be kept below 70% to discourage the growth of insect and plant fungal pathogens. Fertilizer rates and watering must be reduced to discourage fungal pathogens and salt accumulation. The light intensity for the colony should be fairly high. VHO fluorescent bulbs used in sufficient numbers to generate approximately 800-1,000 ft candles at the canopy height are adequate for most plants (i.e. cotton, common bean, lima bean, tomato). Cleanliness is essential in a whitefly colony to maintain optimal rearing conditions.</li>
	<li>Colony Cages: Whiteflies should be maintained on plants in cages, rather than free in the growth room. Cages can be constructed of various materials but must allow for: ventilation, ease of access, ability to prevent whitefly escape or infiltration, and of sufficient size to maintain enough plants to generate the whitefly population needed. Good results have been obtained with organza plus aluminum screening (for strength) as well as whitefly-proof screening (see table of specific equipment and supplies).</li>
	<li>Colony Plant Preparation: The species of plant selected is an important consideration when starting a whitefly colony. The plant should be able to support a high insect population without collapsing. The plant chosen for a nonviruliferous colony should be a nonhost for the virus or viruses that are intended to be transmitted to avoid contamination in the colony and if maintaining a viruliferous colony, it must be host of the virus. The plants should grow fairly quickly to support emergence of new adults within 18 days of being introduced to the colony but not so quickly as to outgrow the cages within four weeks. Dwarf, bush, or patio-type plant cultivars are recommended as they often produce a similar leaf area to taller cultivars but without the stem elongation.</li>
	<li>It is very important to rear plants for the colony in greenhouses in cages that exclude whiteflies and other insects. Use of plants that are infested with whiteflies, thrips, or mites before they enter the colony can cause the colony to collapse. Infested plants also have the potential to be infected with insect-borne viruses which will interfere with virus transmission studies.</li>
</ol>
2. Whitefly Colony Establishment
<ol>
	<li>Start a colony for the first time using clean whiteflies - ones free of plant viruses, other insects, and insect pathogens. These can be obtained from collaborators or from the field. If field collected, whiteflies should be reared for at least 8 weeks on nonhost plants of the plant virus, and check for absence of plant symptoms to be sure they are free of plant viruses.</li>
	<li>Week 1: Introduce clean whiteflies to the first cage of plants. Whiteflies can be introduced by aspiration of known numbers or by gently shaking whiteflies from another source plant depending on the demands on the colony. During week 1, whiteflies are laying eggs on the underside of plant leaf surfaces. An approximate population size can be predicted for each emergence based on the number of adults used to lay eggs and the average number of eggs laid by the female whitefly on the host plant in question.</li>
	<li>Week 2: Begin another cage by introducing whiteflies to new host plants to lay eggs. The plants in the cage started in week 1 will have eggs, and some immature whiteflies covering the undersides of the leaves (Figure 2). Some adult whiteflies introduced during week 1 will still be alive.</li>
	<li>Week 3: Begin a third cage by introducing whiteflies to new plants to lay eggs. The plants in the cage started in week 1 will have immature as well as many new adult whiteflies emerging and there should be a very noticeable increase in the adult whitefly population. The plants in the cage started in week 2 will have many eggs and immature whiteflies covering the undersides of the leaves.</li>
	<li>Week 4: Begin a fourth cage by introducing whiteflies to new plants to lay eggs. The plants in the cage started in week 1 will have many adult whiteflies that are approximately one week post emergence. The plants in the cage started in week 2 will have many new adult whiteflies emerging and there should be a very noticeable increase in whitefly population. The plants in the cage started during week 3 will have many eggs and immature whiteflies covering the undersides of the leaves.</li>
	<li>Weekly after the first 4 weeks: Each week, start a new cage by introducing the adult whiteflies from the fourth oldest cage (the cage started in week 1) onto new plants in a new, clean cage. The old cage should be removed and its plants discarded. These whiteflies are approximately 1 week post-emergence. In the case of the viruliferous colony, these whiteflies will transmit virus to the new plants as well as lay eggs for the next generation of whiteflies.</li>
	<li>Plan transmission experiments to use the whiteflies that will emerge in the third week&#39;s cage. If more whiteflies are needed, plant numbers can be increased and more whiteflies can be added to the week 1 cage to increase whitefly population.</li>
</ol>
3. Method for Inoculation of Test Plants
<ol>
	<li>To insure high transmission rates: 1) Whiteflies must be handled as gently as possible to prevent damage to the insect which will reduce transmission rates; 2) There must be adequate leaf area available for whiteflies to either probe or feed. An increase in acquisition time or an increase in the number of acquisition host plants may increase low rates of infectivity in test plants due to crowding in either the acquisition or inoculation access period.</li>
	<li>Collection of Whiteflies. The following is a procedure for the collection and movement of precise numbers of whiteflies required for experimental purposes. To move large numbers of whiteflies, as is needed for resistance screening for example, it is only necessary to gently shake the whitefly-infested plants over the test plants or the virus-infected acquisition hosts (and later the test plants). Be sure to shake the plants over many locations to minimize aggregation effects.</li>
	<li>Newly emerged adult whiteflies (1-3 days post emergence) are highly active and feed often so tend to give the highest transmission rates 4. Older whiteflies still transmit but at a lower frequency. Multiple whiteflies per test plant (15-40 per plant) should be used for high rates of transmission since a ratio of 1 whitefly per plant often results in unacceptably low transmission rates3,4. The number needed depend upon the virus and the species of the acquisition and test plants.</li>
	<li>Assemble aspiration devices and collection vials. (Figure 3)</li>
	<li>With one hand, hold a yellow plastic card inside the colony cage that contains the whiteflies to be collected. Gently tap the plants to encourage the adult whiteflies to fly. Whiteflies will be attracted to the yellow card and will fly from the plant to the card where they can be collected using an aspirator and a very gentle breath. Never aspirate whiteflies that are feeding on plants. Their stylets are embedded in the plant while feeding so pulling them off the plants will break their stylets and rendering them unable to either acquire or transmit virus (Figure 4).</li>
	<li>Collect about 20 adult whiteflies into a single collection vial to minimize physical damage to the insects from repeated aspirations (Figure 5).</li>
	<li>To change the collection vial, gently tap the vial on a hard surface and cap with Parafilm while the whiteflies are disoriented from the tapping.</li>
	<li>Put a new collect vial on the aspirator and repeat until the number of whiteflies needed are collected. Whiteflies can remain in the collection vials at room temperature for several hours.</li>
	<li>Acquisition. Whiteflies are placed on a virus infected plant and allowed to feed on the infected plants for 48-72 hr. Acquisition periods longer than 72 hr generally do not increase transmission rates. For those viruses transmitted in a nonpersistent or semipersistent manner, 1 hr and several hours, respectively, are sufficient acquisition access periods (Table 1). Best results are obtained when whiteflies are given free access to plants in cages that confine the insects to a single plant or to multiple plants. However some experiments require acquisition from specific leaves in which case clip cages can be used. When using clip cages it is important to accommodate the whitefly feeding preference for the undersides of leaves. Also, crowding of whiteflies in clip cages can reduce transmission rates - acceptable rates of transmission have been achieved using 10 female whiteflies per clip cage of 2.5 cm diameter.</li>
	<li>Inoculation. Prepare a cage appropriate for the size of the inoculation and place the test plants inside. If only one plant is to be inoculated consider a single plant cage. If more than one plant is to be inoculated consider a small PVC&#160;frame/organza bag cage or an aluminum cage (Figure 6). The size of the cage used for inoculation should be slightly larger the size of the plant(s) to be inoculated. Higher transmission rates are obtained when whiteflies are kept close to the plant canopy and not given a lot of free space.</li>
	<li>Collection of whiteflies from acquisition hosts depends on the needs of the experiment. If small numbers are needed, aspirate whiteflies as described above. Place the collection vials of whiteflies inside the cage near the plants and remove the lid (Figure 7). Open the vial and release the whiteflies or invert and gently tap the vial to release the whiteflies. If large numbers are needed simply move acquisition plants with whiteflies into the cages where the test plants are located and gently shake the whiteflies off the acquisition host plant. For both types of introduction, be sure to distribute the whiteflies across the plants to minimize aggregation and insure a uniform inoculation.</li>
	<li>Allow the whiteflies to probe or feed for the appropriate amount of time, again dependent on the virus and to some extent the host plant (Table 1).</li>
	<li>Check the whiteflies at least once during inoculation access period to insure that whiteflies are probing or feeding by opening the cage and turning over some leaves on each plant.</li>
	<li>For the longer inoculation access periods, disturb the plants gently (using a bamboo stick or equivalent) to brush the tops of the plants and encourage the whiteflies to redistribute on the plants. Redistribution of whiteflies helps ensure a higher transmission rate by countering the natural tendency of whiteflies to aggregate.</li>
	<li>Termination. The inoculation access period is ended by killing the whiteflies with approved chemicals. Apply two insecticides one after the other: a contact insecticide to quickly terminate adult whiteflies and a systemic insecticide to terminate any whiteflies that develop in the following weeks and those missed by the contact insecticide (Figure 8). A contact insecticide such as insecticidal soap and systemic insecticides such as imidacloprid or pymetrozine have been used with good success.</li>
</ol>

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	<li>Antignus, Y., Mor, N., Ben Joseph, R., Lapidot, M., Cohen, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=UV+absorbing+plastic+sheets+protect+crops+from+insect+pests+and+from+virus+diseases+vectored+by+insects.">UV absorbing plastic sheets protect crops from insect pests and from virus diseases vectored by insects.</a> Environ. Entomol. 25, 919-924 (1996).</li><li>Cohen, S., Gerling, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epidemiology+of+whitefly-transmitted+viruses.">Epidemiology of whitefly-transmitted viruses.</a> Whiteflies: Their Bionomics, Pest Status and Management. , 211-225 (1990).</li><li>Cohen, S., Antignus, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tomato+yellow+leaf+curl+virus,+a+whitefly-borne+geminivirus+of+tomatoes.">Tomato yellow leaf curl virus, a whitefly-borne geminivirus of tomatoes.</a> Adv. Dis. Vector Res. 10, 259-288 (1994).</li><li>Csizinszky, A., Schuster, D. J., Polston, 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=Effect+of+UV-reflective+mulches+on+tomato+yields+and+on+the+silverleaf+whitefly.">Effect of UV-reflective mulches on tomato yields and on the silverleaf whitefly.</a> HortSci. 34, 911-914 (1999).</li><li>De Barro, P. J., Liu, S. -. S., Boykin, L. M., Dinsdale, A. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bemisia+tabaci:+A+statement+of+species+status.">Bemisia tabaci: A statement of species status.</a> Annu. Rev. Entomol. 56, 1-19 (2011).</li><li>Duffus, J. E., Gerling, D., Mayer, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whitefly-borne+viruses.">Whitefly-borne viruses.</a> Bemisia: 1995 Taxonomy, Biology, Damage, Control and Management. , 255-288 (1996).</li><li>Lapidot, M., Czosnek, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Screening+for+TYLCV-resistant+plants+using+whitefly-mediated+inoculation.">Screening for TYLCV-resistant plants using whitefly-mediated inoculation.</a> Tomato Yellow Leaf Curl Virus Disease. , 329-342 (2007).</li><li>Lapidot, M., Friedmann, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Breeding+for+resistance+to+whitefly-transmitted+geminviruses.">Breeding for resistance to whitefly-transmitted geminviruses.</a> Ann. Appl. Biol. 140, 109-117 (2002).</li><li>Lapidot, M., Polston, J. E., Stansly, P. A., Naranjo, S. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biology+and+epidemiology+of+Bemisia-vectored+viruses.">Biology and epidemiology of Bemisia-vectored viruses.</a> Bemisia: Bionomics and Management of a Global Pest. , 227-231 (2010).</li><li>Navas-Castillo, J., Fiallo-Olive, E., Sanchez-Campos, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Emerging+virus+diseases+transmitted+by+whiteflies.">Emerging virus diseases transmitted by whiteflies.</a> Annu. Rev. Phytopathol. 49, 219-248 (2011).</li><li>Ng, J. C. K., Falk, B. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Virus-vector+interactions+mediating+nonpersistent+and+semipersistent+transmission+of+plant+viruses.">Virus-vector interactions mediating nonpersistent and semipersistent transmission of plant viruses.</a> Ann. Rev. Phytopathol. 44, 212-21 (2006).</li><li>Polston, J. E., Cohen, L., Sherwood, T. A., Ben-Joseph, R., Lapidot, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Capsicum+species:+Symptomless+hosts+and+reservoirs+of+Tomato+yellow+leaf+curl+virus+(TYLCV).">Capsicum species: Symptomless hosts and reservoirs of Tomato yellow leaf curl virus (TYLCV).</a> Phytopathology. 96, 447-452 (2006).</li><li>Polston, J. E., Hiebert, E., McGovern, R. J., Stansly, P. A., Schuster, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Host+range+of+Tomato+mottle+virus,+a+new+geminivirus+infecting+tomato+in+Florida.">Host range of Tomato mottle virus, a new geminivirus infecting tomato in Florida.</a> Plant Dis. 77, 1181-1184 (1993).</li><li>Polston, J. E., Sherwood, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pymetrozine+interferes+with+transmission+of+Tomato+yellow+leaf+curl+virus+by+the+whitefly+Bemisia+tabaci+(Hemiptera:+Aleyrodidae.">Pymetrozine interferes with transmission of Tomato yellow leaf curl virus by the whitefly Bemisia tabaci (Hemiptera: Aleyrodidae.</a> Phytoparasitica. 31, 490-498 (2003).</li><li>Schuster, D. J., Thompson, S., Ortega, L. D., Polston, 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=Laboratory+evaluation+of+products+to+reduce+settling+of+sweetpotato+whitefly+adults.">Laboratory evaluation of products to reduce settling of sweetpotato whitefly adults.</a> J. Econ. Ento. 102, 1482-1489 (2009).</li><li>King, A. M. Q., Adams, M. J., Carstons, E. B., Lefkowitz, E. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Virus Taxonomy, Ninth Report of the International Committee on Taxonomy of Viruses. , (2012).</li><li>Wintermantel, W. M., Stansly, P. A., Naranjo, S. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transmission+efficiency+and+epidemiology+of+Criniviruses.">Transmission efficiency and epidemiology of Criniviruses.</a> Bemisia: Bionomics and Management of a Global Pest. , 319-332 (2010).</li><li>Yang, Y., Sherwood, T. A., Patte, C. P., Hiebert, E., Polston, 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=Use+of+Tomato+yellow+leaf+curl+virus+(TYLCV)+Rep+gene+sequences+to+engineer+TYLCV+resistance+in+tomato.">Use of Tomato yellow leaf curl virus (TYLCV) Rep gene sequences to engineer TYLCV resistance in tomato.</a> Phytopathology. 94, 490-496 (2004).</li><li>Zehnder, G. W., Yao, C., Murphy, J. F., Sikora, E. R., Kloepper, J. W., Schuster, D. J., Polston, 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=Microbe-Induced+Resistance+Against+Pathogens+and+Herbivores:+Evidence+of+Effectiveness+in+Agriculture.">Microbe-Induced Resistance Against Pathogens and Herbivores: Evidence of Effectiveness in Agriculture.</a> Induced Plant Defenses Against Pathogens. , 335-355 (1999).</li></ol>

No conflicts of interest declared.

The methods described here were developed over a period of two decades and are based on the basic information provided by many studies of whitefly transmission, behavior, and biology. Since there are many publications, this manuscript refers primarily to reviews and to a few selected specific publications to illustrate the type of data used to develop this technique. It is very important in this procedure that the whiteflies are handled gently since breaking of legs and antennas can lead to abnormal behaviors and lower rates of transmission. Collection with vacuums and other mechanical suction devices has never produced good results. Another consideration for success is that the number of whiteflies used to obtain 100% transmission to test plants, must be determined for each combination of plant host and virus. This optimal ratio only needs to be established once. Best results have been obtained using young adult whiteflies (1-3 days past emergence) since they give the highest rates of transmission. Older adults can be used but larger numbers of whiteflies will be needed to compensate for their reduced rates of transmission. Females are known to transmit at higher rates than males, since they feed more often than males. However it is usually not worth the time to separate the sexes for transmission. This procedure can be modified in many ways to suit the resources and needs of the researcher.
When working with multiple viruses it is best to keep a single colony of whiteflies reared on nonhost plants of the virus. These whiteflies can be collected placed on virus-infected plants for acquisition and then after acquisition the whiteflies can be moved again to the test plants. Plants intended for the colony should be grown in greenhouses in cages to prevent the introduction of whiteflies that might infect them with another virus. Infestation of these plants with mites, thrips, and other plant pests can compromise the health of the colony and cause it to collapse. Finally watering of the colony plants has to be done with care. These plants are subject to extreme stress by the feeding of so many whiteflies and the lower than normal light conditions. Root rotting fungi which are introduced on the peat in the soilless mixes can become a problem if the plants are overwatered even once. Pretreatment of the plants with a fungicide drench can eliminate some of the problems.
While whitefly transmission of plant viruses can be time consuming and require valuable resources (such as growth rooms) it is essential for the transmission of some viruses for which we have no other means of transmission. It is also a valuable means of screening plants for resistance to viruses, as it uses the same type of transmission that plants in the field will be expected to resist. The use of more artificial means of transmission do not always yield the best results in evaluations of germplasm for virus-resistance.

<table border="1">
	<tbody>
		<tr>
			<td>Name of the Item</td>
			<td>Company</td>
			<td>Catalogue No.</td>
			<td>Comments (optional)</td>
		</tr>
		<tr>
			<td>Whitefly proof screening</td>
			<td>Klayman Meteor Ltd. Petah Tikva 49130, Israel</td>
			<td>---</td>
			<td>BioNet 50 mesh screen Antivirus Insect Screen</td>
		</tr>
		<tr>
			<td>Whitefly proof screening</td>
			<td>Hummert International (www.hummert.com)</td>
			<td>27-4050-1 - 27-4062-1</td>
			<td>BioNet 50 mesh screen Antivirus Insect Screen</td>
		</tr>
		<tr>
			<td>Whitefly Collection Devices</td>
			<td>BioQuip</td>
			<td>1135A</td>
			<td>Can also be made using eye droppers, flexible plastic tubing, plastic vacuum adaptors, and cheesecloth</td>
		</tr>
		<tr>
			<td>Collection vials</td>
			<td>BioQuip</td>
			<td>8909</td>
			<td>Vials used depend on which aspirator is used.</td>
		</tr>
		<tr>
			<td>Yellow plastic cards</td>
			<td>any company</td>
			<td>---</td>
			<td>Any bright yellow plastic card will work, can be purchased in many places.</td>
		</tr>
		<tr>
			<td>Whitefly Cages</td>
			<td>---</td>
			<td>---</td>
			<td>These are not commercially available and must be constructed. 60 cm x 60 cm x 60 cm cages work well for us. Our preference is for those constructed of aluminum window screening, nylon organdy, vinyl plastic, with a structure created using aluminum window frames. Doors are attached using piano hinges and sealed with weather stripping; tops of the cages are vinyl plastic. Cages can also be made of 1) plexiglass glued together on the edges with ventilation provided by screened openings cut on the side pieces; 2) large bags of nylon organdy, organza, or 50 mesh whitefly screening with internal supports provided by PVC pipes and fittings cut to desired lengths and assembled inside the bags, 3) wood frames and nylon organdy, organza, or 50 mesh whitefly screening.</td>
		</tr>
	</tbody>
</table>

transmitting plant viruses using whiteflies

Whiteflies, Hemiptera: Aleyrodidae, Bemisia tabaci, a complex of morphologically indistinquishable species5, are vectors of many plant viruses. Several genera of these whitefly-transmitted plant viruses (Begomovirus, Carlavirus, Crinivirus, Ipomovirus, Torradovirus) include several hundred species of emerging and economically significant pathogens of important food and fiber crops (reviewed by9,10,16). These viruses do not replicate in their vector but nevertheless are moved readily from plant to plant by the adult whitefly by various means (reviewed by2,6,7,9,10,11,17). For most of these viruses whitefly feeding is required for acquisition and inoculation, while for others only probing is required. Many of these viruses are unable or cannot be easily transmitted by other means. Therefore maintenance of virus cultures, biological and molecular characterization (identification of host range and symptoms)3,13, ecology2,12, require that the viruses be transmitted to experimental hosts using the whitefly vector. In addition the development of new approaches to management, such as evaluation of new chemicals14 or compounds15, new cultural approaches1,4,19, or the selection and development of resistant cultivars7,8,18, requires the use of whiteflies for virus transmission. The use of whitefly transmission of plant viruses for the selection and development of resistant cultivars in breeding programs is particularly challenging7. Effective selection and screening for resistance employs large numbers of plants and there is a need for 100% of the plants to be inoculated in order to find the few genotypes which possess resistance genes. These studies use very large numbers of viruliferous whiteflies, often several times per year.
Whitefly maintenance described here can generate hundreds or thousands of adult whiteflies on plants each week, year round, without the contamination of other plant viruses. Plants free of both whiteflies and virus must be produced to introduce into the whitefly colony each week. Whitefly cultures must be kept free of whitefly pathogens, parasites, and parasitoids that can reduce whitefly populations and/or reduce the transmission efficiency of the virus. Colonies produced in the manner described can be quickly scaled to increase or decrease population numbers as needed, and can be adjusted to accommodate the feeding preferences of the whitefly based on the plant host of the virus.
There are two basic types of whitefly colonies that can be maintained: a nonviruliferous and a viruliferous whitefly colony. The nonviruliferous colony is composed of whiteflies reared on virus-free plants and allows the weekly availability of whiteflies which can be used to transmit viruses from different cultures. The viruliferous whitefly colony, composed of whiteflies reared on virus-infected plants, allows weekly availability of whiteflies which have acquired the virus thus omitting one step in the virus transmission process.

These methods for whitefly colony establishment and maintenance, as well as manipulation to transmit plant viruses have been used successfully in a number of studies4,11,12,14 as well as many others not cited. Using these methods with persistently transmitted viruses, we have routinely obtained the desired rates of transmission of 100% for selection for resistance, screening insecticides for their ability to interfere with transmission (and evaluation of resistance inducing compounds)13,14,17,18 (Figure 9). The procedures described here can and have been be adapted to many locations in both public and private research facilities.
<table border="1">
	<tbody>
		<tr>
			<td>Type of Transmission</td>
			<td>Acquisition Access Period (hour)</td>
			<td>Inoculation Access Period (hour)</td>
			<td>Ref.</td>
		</tr>
		<tr>
			<td>Nonpersistent</td>
			<td>1</td>
			<td>2-24</td>
			<td>10</td>
		</tr>
		<tr>
			<td>Semipersistent</td>
			<td>6-24</td>
			<td>8-24</td>
			<td>10, 16</td>
		</tr>
		<tr>
			<td>Persistent</td>
			<td>48-72</td>
			<td>48-72</td>
			<td>3, 6, 8, 9</td>
		</tr>
	</tbody>
</table>
Table 1. Estimated times that can be used to produce high rates of transmission of viruses with different types of relationships with their vector Bemisia tabaci species complex. Estimated times are based on published minimum times required for 80-100% transmission and modified in some cases to account for latent periods and differences among whitefly populations, virus, and host plants.
<img alt="Figure 1" src="/files/ftp_upload/4332/4332fig1.jpg" /> 
Figure 1. An example of a viruliferous whitefly colony. This growth room contains cages of plants infected with a begomovirus and infested with viruliferous whiteflies.
<img alt="Figure 2" src="/files/ftp_upload/4332/4332fig2.jpg" /> 
Figure 2. Left: Immatures stages of the whitefly, Bemisia tabaci MEAM clade sensu De Barro6 approximately 1 week post-emergence and Right: an adult whitefly, an effective vector of many plant viruses.
<img alt="Figure 3" fo:content-width="5.5in" fo:src="/files/ftp_upload/4332/4332fig3highres.jpg" src="/files/ftp_upload/4332/4332fig3.jpg" /> 
Figure 3. Types of collection devices that can be used to collect whiteflies for plant virus transmission.
<img alt="Figure 4" src="/files/ftp_upload/4332/4332fig4.jpg" /> 
Figure 4. Left: Yellow plastic card used to collect whitefly adults showing whiteflies ready for collection. Right: Collecting whiteflies from a yellow plastic card. Whiteflies can be seen on the card as small white/gray spots.
<img alt="Figure 5" src="/files/ftp_upload/4332/4332fig5.jpg" /> 
Figure 5. Whiteflies collected by aspiration (20 in a vial) from the colony ready for either acquisition or inoculation.
<img alt="Figure 6" fo:content-width="5.5in" fo:src="/files/ftp_upload/4332/4332fig6highres.jpg" src="/files/ftp_upload/4332/4332fig6.jpg" /> 
Figure 6. Upper Left: Single plant cages used for either acquisition or inoculation. Upper Right: Larger cages made of pvc pipe and organza. Lower: Aluminum/screen and organdy cages in a greenhouse containing plants intended for a whitefly colony.
<img alt="Figure 7" src="/files/ftp_upload/4332/4332fig7.jpg" /> 
Figure 7. Whiteflies being transferred to a single plant for inoculation (or acquisition).
<img alt="Figure 8" src="/files/ftp_upload/4332/4332fig8.jpg" /> 
Figure 8. Addition of a systemic insecticide drench to terminate the whitefly inoculation access period.
<img alt="Figure 9" src="/files/ftp_upload/4332/4332fig9.jpg" /> 
Figure 9. Field plants under evaluation for resistance to Tomato yellow leaf curl virus. All plants were inoculated as 5 week old seedlings and then transplanted to the field for evaluation. Plants in the foreground are a resistant germplasm line, plants in the background are susceptible germplasm line.

Watch this Scientific Journal Video about Transmitting Plant Viruses Using Whiteflies at JoVE.com

Transmitting Plant Viruses Using Whiteflies

Improving our understanding of and our ability to manage many of the insect-transmitted plant viruses requires the use of the vector. Insect transmission of plant viruses is a tritrophic interaction, and as such requires the manipulation of insects, virus, and plant. Vectors must be reared in large numbers and manipulated in such a way as to insure high rates of transmission to test plants. The basics of rearing and manipulating the whitefly, Bemisa tabaci (Hemiptera: Aleyrodidae), a vector of plant viruses belonging to several emerging plant virus genera, representing a large number of economically significant viruses is presented.

Whiteflies, Hemiptera: Aleyrodidae, Bemisia tabaci, a complex of morphologically indistinquishable species5, are vectors of many plant viruses. Several ...

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28.7K Views.  University of Florida. Improving our understanding of and our ability to manage many of the insect-transmitted plant viruses requires the use of the vector. Insect transmission of plant viruses is a tritrophic interaction, and as such requires the manipulation of insects, virus, and plant. Vectors must be reared in large numbers and manipulated in such a way as to insure high rates of transmission to test plants. The basics of rearing and manipulating the whitefly, Bemisa tabaci (H...

Video: Transmitting Plant Viruses Using Whiteflies

Testing the Physiological Barriers to Viral Transmission in Aphids Using Microinjection

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

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

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

Testing Nicotine Tolerance in Aphids Using an Artificial Diet Experiment

Plants may upregulate the production of many different seconday metabolites in response to insect feeding. One of these metabolites, nicotine, is well know to have insecticidal properties. One response of tobacco plants to herbivory, or being gnawed upon by insects, is to increase the production of this neurotoxic alkaloid. Here, we will demonstrate how to set up an experiment to address this question of whether a tobacco-adapted strain of the green peach aphid, Myzus persicae, can tolerate higher levels of nicotine than the a strain of this insect that does not infest tobacco in the field.

In this video and assay is demonstrated that tests the tolerance of nicotine to two types of aphids one that infests tobacco plants in the field and one that does not.

Plants may upregulate the production of many different seconday metabolites in response to insect feeding. One of these metabolites, nicotine, is well ...

Virus-induced Gene Silencing (VIGS) in Nicotiana benthamiana and Tomato

RNA interference (RNAi) is a highly specific gene-silencing phenomenon triggered by dsRNA1. This silencing mechanism uses two major classes of RNA regulators: microRNAs, which are produced from non-protein coding genes and short interfering RNAs (siRNAs). Plants use RNAi to control transposons and to exert tight control over developmental processes such as flower organ formation and leaf development2,3,4. Plants also use RNAi to defend themselves against infection by viruses. Consequently, many viruses have evolved suppressors of gene silencing to allow their successful colonization of their host5.
Virus-induced gene silencing (VIGS) is a method that takes advantage of the plant RNAi-mediated antiviral defense mechanism. In plants infected with unmodified viruses the mechanism is specifically targeted against the viral genome. However, with virus vectors carrying sequences derived from host genes, the process can be additionally targeted against the corresponding host mRNAs. VIGS has been adapted for high-throughput functional genomics in plants by using the plant pathogen Agrobacterium tumefaciens to deliver, via its Ti plasmid, a recombinant virus carrying the entire or part of the gene sequence targeted for silencing. Systemic virus spread and the endogenous plant RNAi machinery take care of the rest. dsRNAs corresponding to the target gene are produced and then cleaved by the ribonuclease Dicer into siRNAs of 21 to 24 nucleotides in length. These siRNAs ultimately guide the RNA-induced silencing complex (RISC) to degrade the target transcript2.
Different vectors have been employed in VIGS and one of the most frequently used is based on tobacco rattle virus (TRV). TRV is a bipartite virus and, as such, two different A. tumefaciens strains are used for VIGS. One carries pTRV1, which encodes the replication and movement viral functions while the other, pTRV2, harbors the coat protein and the sequence used for VIGS6,7. Inoculation of Nicotiana benthamiana and tomato seedlings with a mixture of both strains results in gene silencing. Silencing of the endogenous phytoene desaturase (PDS) gene, which causes photobleaching, is used as a control for VIGS efficiency. It should be noted, however, that silencing in tomato is usually less efficient than in N. benthamiana. RNA transcript abundance of the gene of interest should always be measured to ensure that the target gene has efficiently been down-regulated. Nevertheless, heterologous gene sequences from N. benthamiana can be used to silence their respective orthologs in tomato and vice versa8.

Virus-induced Gene Silencing VIGS in Nicotiana benthamiana and Tomato

Description of a virus-induced gene silencing (VIGS) method for knock-down of gene expression in Nicotiana benthamiana and tomato.

RNA interference (RNAi) is a highly specific gene-silencing phenomenon triggered by dsRNA1. This silencing mechanism uses two major classes of RNA ...

Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

Light mediates an array of developmental and adaptive processes throughout the life cycle of a plant. Plants utilize light-absorbing molecules called photoreceptors to sense and adapt to light. The red/far-red light-absorbing phytochrome photoreceptors have been studied extensively. Phytochromes exist as a family of proteins with distinct and overlapping functions in all higher plant systems in which they have been studied1. Phytochrome-mediated light responses, which range from seed germination through flowering and senescence, are often localized to specific plant tissues or organs2. Despite the discovery and elucidation of individual and redundant phytochrome functions through mutational analyses, conclusive reports on distinct sites of photoperception and the molecular mechanisms of localized pools of phytochromes that mediate spatial-specific phytochrome responses are limited. We designed experiments based on the hypotheses that specific sites of phytochrome photoperception regulate tissue- and organ-specific aspects of photomorphogenesis, and that localized phytochrome pools engage distinct subsets of downstream target genes in cell-to-cell signaling. We developed a biochemical approach to selectively reduce functional phytochromes in an organ- or tissue-specific manner within transgenic plants. Our studies are based on a bipartite enhancer-trap approach that results in transactivation of the expression of a gene under control of the Upstream Activation Sequence (UAS) element by the transcriptional activator GAL43. The biliverdin reductase (BVR) gene under the control of the UAS is silently maintained in the absence of GAL4 transactivation in the UAS-BVR parent4. Genetic crosses between a UAS-BVR transgenic line and a GAL4-GFP enhancer trap line result in specific expression of the BVR gene in cells marked by GFP expression4. BVR accumulation in Arabidopsis plants results in phytochrome chromophore deficiency in planta5-7. Thus, transgenic plants that we have produced exhibit GAL4-dependent activation of the BVR gene, resulting in the biochemical inactivation of phytochrome, as well as GAL4-dependent GFP expression. Photobiological and molecular genetic analyses of BVR transgenic lines are yielding insight into tissue- and organ-specific phytochrome-mediated responses that have been associated with corresponding sites of photoperception4, 7, 8. Fluorescence Activated Cell Sorting (FACS) of GFP-positive, enhancer-trap-induced BVR-expressing plant protoplasts coupled with cell-type-specific gene expression profiling through microarray analysis is being used to identify putative downstream target genes involved in mediating spatial-specific phytochrome responses. This research is expanding our understanding of sites of light perception, the mechanisms through which various tissues or organs cooperate in light-regulated plant growth and development, and advancing the molecular dissection of complex phytochrome-mediated cell-to-cell signaling cascades.

Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

The molecular basis of spatial-specific phytochrome responses is being investigated using transgenic plants that exhibit tissue- and organ-specific phytochrome deficiencies. The isolation of specific cells exhibiting induced phytochrome chromophore depletion by Fluorescence-Activated Cell Sorting followed by microarray analyses is being utilized to identify genes involved in spatial-specific phytochrome responses.

Light mediates an array of developmental and adaptive processes throughout the life cycle of a plant. Plants utilize light-absorbing molecules called ...

Profiling Thiol Redox Proteome Using Isotope Tagging Mass Spectrometry

Pseudomonas syringae pv. tomato strain DC3000 not only causes bacterial speck disease in Solanum lycopersicum but also on Brassica species, as well as on Arabidopsis thaliana, a genetically tractable host plant1,2. The accumulation of reactive oxygen species (ROS) in cotyledons inoculated with DC3000 indicates a role of ROS in modulating necrotic cell death during bacterial speck disease of tomato3. Hydrogen peroxide, a component of ROS, is produced after inoculation of tomato plants with Pseudomonas3. Hydrogen peroxide can be detected using a histochemical stain 3'-3' diaminobenzidine (DAB)4. DAB staining reacts with hydrogen peroxide to produce a brown stain on the leaf tissue4. ROS has a regulatory role of the cellular redox environment, which can change the redox status of certain proteins5. Cysteine is an important amino acid sensitive to redox changes. Under mild oxidation, reversible oxidation of cysteine sulfhydryl groups serves as redox sensors and signal transducers that regulate a variety of physiological processes6,7. Tandem mass tag (TMT) reagents enable concurrent identification and multiplexed quantitation of proteins in different samples using tandem mass spectrometry8,9. The cysteine-reactive TMT (cysTMT) reagents enable selective labeling and relative quantitation of cysteine-containing peptides from up to six biological samples. Each isobaric cysTMT tag has the same nominal parent mass and is composed of a sulfhydryl-reactive group, a MS-neutral spacer arm and an MS/MS reporter10. After labeling, the samples were subject to protease digestion. The cysteine-labeled peptides were enriched using a resin containing anti-TMT antibody. During MS/MS analysis, a series of reporter ions (i.e., 126-131 Da) emerge in the low mass region, providing information on relative quantitation. The workflow is effective for reducing sample complexity, improving dynamic range and studying cysteine modifications. Here we present redox proteomic analysis of the Pst DC3000 treated tomato (Rio Grande) leaves using cysTMT technology. This high-throughput method has the potential to be applied to studying other redox-regulated physiological processes.

Reactive oxygen species level is elevated when cells encounter stress conditions. Here we show the example of 3'-3' diaminobenzidine staining as well as cysTMT labeling and mass spectrometry to profile the redox proteome in Pseudomonas syringae treated tomato leaves.

Pseudomonas syringae pv. tomato strain DC3000 not only causes bacterial speck disease in Solanum lycopersicum but also on Brassica species, as well as on ...

Fruit Volatile Analysis Using an Electronic Nose

Numerous and diverse physiological changes occur during fruit ripening, including the development of a specific volatile blend that characterizes fruit aroma. Maturity at harvest is one of the key factors influencing the flavor quality of fruits and vegetables1. The validation of robust methods that rapidly assess fruit maturity and aroma quality would allow improved management of advanced breeding programs, production practices and postharvest handling. 
Over the last three decades, much research has been conducted to develop so-called electronic noses, which are devices able to rapidly detect odors and flavors2-4. Currently there are several commercially available electronic noses able to perform volatile analysis, based on different technologies. The electronic nose used in our work (zNose, EST, Newbury Park, CA, USA), consists of ultra-fast gas chromatography coupled with a surface acoustic wave sensor (UFGC-SAW). This technology has already been tested for its ability to monitor quality of various commodities, including detection of deterioration in apple5; ripeness and rot evaluation in mango6; aroma profiling of thymus species7; C6 volatile compounds in grape berries8; characterization of vegetable oil9 and detection of adulterants in virgin coconut oil10. 
This system can perform the three major steps of aroma analysis: headspace sampling, separation of volatile compounds, and detection. In about one minute, the output, a chromatogram, is produced and, after a purging cycle, the instrument is ready for further analysis. The results obtained with the zNose can be compared to those of other gas-chromatographic systems by calculation of Kovats Indices (KI). Once the instrument has been tuned with an alkane standard solution, the retention times are automatically converted into KIs. However, slight changes in temperature and flow rate are expected to occur over time, causing retention times to drift. Also, depending on the polarity of the column stationary phase, the reproducibility of KI calculations can vary by several index units11. A series of programs and graphical interfaces were therefore developed to compare calculated KIs among samples in a semi-automated fashion. These programs reduce the time required for chromatogram analysis of large data sets and minimize the potential for misinterpretation of the data when chromatograms are not perfectly aligned.
We present a method for rapid volatile compound analysis in fruit. Sample preparation, data acquisition and handling procedures are also discussed.

A rapid method for volatile compound analysis in fruit is described. The volatile compounds present in the headspace of a homogenate of the sample are rapidly separated and detected with ultra-fast gas chromatography (GC) coupled with a surface acoustic wave (SAW) sensor. A procedure for data handling and analysis is also discussed.

Numerous and diverse physiological changes occur during fruit ripening, including the development of a specific volatile blend that characterizes fruit ...

Glycan Profiling of Plant Cell Wall Polymers using Microarrays

Plant cell walls are complex matrixes of heterogeneous glycans which play an important role in the physiology and development of plants and provide the raw materials for human societies (e.g. wood, paper, textile and biofuel industries)1,2. However, understanding the biosynthesis and function of these components remains challenging.
Cell wall glycans are chemically and conformationally diverse due to the complexity of their building blocks, the glycosyl residues. These form linkages at multiple positions and differ in ring structure, isomeric or anomeric configuration, and in addition, are substituted with an array of non-sugar residues. Glycan composition varies in different cell and/or tissue types or even sub-domains of a single cell wall3. Furthermore, their composition is also modified during development1, or in response to environmental cues4.
In excess of 2,000 genes have Plant cell walls are complex matrixes of heterogeneous glycans been predicted to be involved in cell wall glycan biosynthesis and modification in Arabidopsis5. However, relatively few of the biosynthetic genes have been functionally characterized 4,5. Reverse genetics approaches are difficult because the genes are often differentially expressed, often at low levels, between cell types6. Also, mutant studies are often hindered by gene redundancy or compensatory mechanisms to ensure appropriate cell wall function is maintained7. Thus novel approaches are needed to rapidly characterise the diverse range of glycan structures and to facilitate functional genomics approaches to understanding cell wall biosynthesis and modification.
Monoclonal antibodies (mAbs)8,9 have emerged as an important tool for determining glycan structure and distribution in plants. These recognise distinct epitopes present within major classes of plant cell wall glycans, including pectins, xyloglucans, xylans, mannans, glucans and arabinogalactans. Recently their use has been extended to large-scale screening experiments to determine the relative abundance of glycans in a broad range of plant and tissue types simultaneously9,10,11.
Here we present a microarray-based glycan screening method called Comprehensive Microarray Polymer Profiling (CoMPP) (Figures 1 &amp; 2)10,11 that enables multiple samples (100 sec) to be screened using a miniaturised microarray platform with reduced reagent and sample volumes. The spot signals on the microarray can be formally quantified to give semi-quantitative data about glycan epitope occurrence. This approach is well suited to tracking glycan changes in complex biological systems12 and providing a global overview of cell wall composition particularly when prior knowledge of this is unavailable.

A technique called Comprehensive Microarray Polymer Profiling (CoMPP) for the characterisation of plant cell wall glycans is described. This method combines the specificity of monoclonal antibodies directed to defined glycan-epitopes with a miniature microarray analytical platform allowing screening of glycan occurrence in a broad range of biological contexts.

Plant cell walls are complex matrixes of heterogeneous glycans which play an important role in the physiology and development of plants and provide the ...

High-throughput Titration of Luciferase-expressing Recombinant Viruses

Standard plaque assays to determine infectious viral titers can be time consuming, are not amenable to a high volume of samples, and cannot be done with viruses that do not form plaques. As an alternative to plaque assays, we have developed a high-throughput titration method that allows for the simultaneous titration of a high volume of samples in a single day. This approach involves infection of the samples with a Firefly luciferase tagged virus, transfer of the infected samples onto an appropriate permissive cell line, subsequent addition of luciferin, reading of plates in order to obtain luminescence readings, and finally the conversion from luminescence to viral titers. The assessment of cytotoxicity using a metabolic viability dye can be easily incorporated in the workflow in parallel and provide valuable information in the context of a drug screen. This technique provides a reliable, high-throughput method to determine viral titers as an alternative to a standard plaque assay.

This article presents a high-throughput luciferase expression-based method of titrating various RNA and DNA viruses using automated and manual liquid handlers.

Standard plaque assays to determine infectious viral titers can be time consuming, are not amenable to a high volume of samples, and cannot be done with ...

A Multi-detection Assay for Malaria Transmitting Mosquitoes

The Anopheles gambiae species complex includes the major malaria transmitting mosquitoes in Africa. Because these species are of such medical importance, several traits are typically characterized using molecular assays to aid in epidemiological studies. These traits include species identification, insecticide resistance, parasite infection status, and host preference. Since populations of the Anopheles gambiae complex are morphologically indistinguishable, a polymerase chain reaction (PCR) is traditionally used to identify species. Once the species is known, several downstream assays are routinely performed to elucidate further characteristics. For instance, mutations known as KDR in a para gene confer resistance against DDT and pyrethroid insecticides. Additionally, enzyme-linked immunosorbent assays (ELISAs) or Plasmodium parasite DNA detection PCR assays are used to detect parasites present in mosquito tissues. Lastly, a combination of PCR and restriction enzyme digests can be used to elucidate host preference (e.g., human vs. animal blood) by screening the mosquito bloodmeal for host-specific DNA. We have developed a multi-detection assay (MDA) that combines all of the aforementioned assays into a single multiplex reaction genotyping 33SNPs for 96 or 384 samples at a time. Because the MDA includes multiple markers for species, Plasmodium detection, and host blood identification, the likelihood of generating false positives or negatives is greatly reduced from previous assays that include only one marker per trait. This robust and simple assay can detect these key mosquito traits cost-effectively and in a fraction of the time of existing assays.

Malaria transmitting mosquitoes have a number of epidemiologically important characteristics that can only be detected using molecular techniques. Utilizing a MALDI-TOF based SNP genotyping platform, we developed an assay for simultaneously detecting multiple key traits (species, insecticide resistance, parasite infection and host choice) of malaria vectors.

The Anopheles gambiae species complex includes the major malaria transmitting mosquitoes in Africa. Because these species are of such medical importance, ...

Imaging Spatial Reorganization of a MAPK Signaling Pathway Using the Tobacco Transient Expression System

Visualization of dynamic signaling events in live cells has been a challenge. We expanded an established transient expression system, the biomolecular fluorescent complementation (BiFC) assay in tobacco epidermal cells, from testing protein-protein interaction to monitoring spatial distribution of signal transduction in plant cells. In this protocol, we used the BiFC assay to show that the interaction and the signaling between the Arabidopsis MAPKKK YODA and MAPK6 occur at the plasma membrane. When the scaffold protein BASL was co-expressed, the YODA-MAPK interaction redistributed and spatially co-polarized with CFP-BASL. This modified tobacco expression system allows for quick examination of signaling localization and dynamic changes (less than 4 days) and can accommodate multiple of fluorescent protein colors (at least three). We also presented detailed methods to quantify protein distribution (asymmetric spatial localization, or &#34;polarization&#34;) in tobacco cells. This advanced tobacco expression system has a potential to be used widely for quick testing of dynamic signaling events in live plant cells.

At the subcellular level, signaling events are dynamically modulated by developmental and environmental cues. Here we describe a protocol that employs the tobacco transient expression system to monitor dynamic protein-protein interaction and to disclose spatial organization of signal transduction in plant cells.

Visualization of dynamic signaling events in live cells has been a challenge. We expanded an established transient expression system, the biomolecular ...