We thank D. Ashton, V. Barkley, K. Beimfohr, F. Bojorquez, J. Cantrell, G. Castro, R. Christensen, J. Fearn, C. Jara, D. Meade, G. Owens, L. Rodarte, D. Sieglaff, A. Sonoqui, M. Stefanek, B. Stuart, J. Trejo, A. Slade and E. Yescas for technical assistance. Partial support was provided by USDA-Agricultural Research Service, USDA-National Institute for Food and Agriculture Extension IPM Program and Pest Management Alternatives Special Projects, Cotton Incorporated, Arizona Cotton Growers Association, Cotton Foundations, USDA-CREES, NAPIAP (Western Region), and Western Region IPM Special Projects.

NOTE: The techniques described below are considered partial life tables because they do not explicitly include reproduction or mortality of the adult stages. The term cohort is equivalent to generation because it examines mortality from the egg to the adult stage.
1. Establish Field Sites
<ol>
	<li>Conduct life tables at any time during the growth of the crop once insects are present. The choice of when to initiate studies will depend on the goals and objectives of the research.</li>
	<li>Select two rows of crop near the center of the plot to minimize edge effects from surrounding plots or uncultivated areas. Mark the head of each row with a wire flag or a wooden stake to facilitate easy relocation once the crop gets large.</li>
	<li>Move 3-4 m in from edge of plot parallel to the row. Use one row to establish egg cohorts and the second to establish nymph cohorts</li>
</ol>
2. Establish Egg Cohorts
<ol>
	<li>Use an 8X hand lens to search for newly laid eggs on the underside of leaves near the top of the main stem of the cotton plant (generally the second or third node from the top). 
	NOTE: Life stages of B. tabaci are generally distributed vertically in the plant canopy with eggs near the top of the plant and progressively older nymphal stages below. This results because 1st instar nymphs settle on the same leaf as the egg and the plant adds leaves above the initial oviposition site as it grows.
	<ol>
		<li>Use a higher power 15X lens to observe fresh eggs and verify identification before using the 8x lens to mark the insect. Fresh eggs have a bright white coloration under the lens and stand out from eggs that are older (Figures 1A and 1B). Eggs darken to a tannish color as they mature.</li>
	</ol>
	</li>
	<li>Use a non-toxic, ultra-fine-point black permanent marker to draw a small circle around the egg. Draw the circle small enough to minimize the chance of a female laying another egg within the circle later on.
	<ol>
		<li>Cut a hole or slot in the side of the 8X hand lens with a hacksaw or drill bit so that the pen can be inserted and viewed through the lens when drawing (Figure 2A).</li>
	</ol>
	</li>
	<li>Repeat this process on the same leaf, if possible, to mark other eggs, marking a total of no more than four eggs on a single leaf and no more than one egg per leaf sector. For cotton, the leaf is subdivided into four sectors by three major leaf veins (Figure 3).</li>
	<li>Tie a small lightweight cardboard tag around the petiole of the leaf containing marked egg(s). Number the tag and include notation for the plot or treatment number depending on the experimental design (Figure 2B).</li>
	<li>Tie a 1 m long length of flagging tape around the main stem near the top of the plant. Use a bow-style tie so that the tape can be easily relocated as necessary to keep it visible for repeated visits to this location in the field.</li>
	<li>Record leaf number and positional information on a portable analog or electronic notebook (Table 1). Positional information uses the sectors of the leaf to more finely note location (e.g., 1-1, 1-2, 1-4 denote eggs in sectors 1, 2 and 4 on leaf #1).</li>
	<li>Establish each cohort on a single day. 
	NOTE: A cohort in a given plot is comprised of a minimum of 50 eggs total.
	<ol>
		<li>Use no more than one leaf per plant to better distribute the members of the cohort. 
		NOTE: Depending on insect density, this may comprise anywhere from 13 leaves with 3-4 eggs each or 50 leaves with one egg each. The individual plants are selected to distribute the marked leaves along as much of the row as possible.</li>
	</ol>
	</li>
</ol>
3. Establish Nymph Cohorts
<ol>
	<li>Use an 8X hand lens to search for newly settled 1st instar nymphs on the underside of leaves about 3-5 leaves down from the top of the main stem of the cotton plant. Use a 15X lens to verify identification before marking using the 8X lens.</li>
	<li>Use a non-toxic, ultra-fine-point black permanent marker to draw a small circle around the nymph. Draw the circle small enough to minimize the chance of a crawler settling within the circle later on. 
	NOTE: Newly hatched 1st instar nymphs are called crawlers, which can move several cm during the first few hours after egg hatch. It then &#34;settles&#34; onto a site where it will feed and molt without ever moving again until the adult emerges. These settled 1st instar nymphs (Figure 1C) are distinctive from the crawlers. First, they are immobile and second, they are more 2-dimensional and lay tight and flat on the leaf and have a slightly more translucent amber color.
	<ol>
		<li>Cut a hole or slot in the side of the 8X hand lens with a hacksaw or drill bit so that the pen can be inserted and viewed through the lens when drawing (Figure 2A).</li>
	</ol>
	</li>
	<li>Repeat this process on the same leaf, if possible, to mark other nymphs, marking no more than four nymphs on a single leaf and no more than one nymph per leaf sector. For cotton, the leaf is subdivided into four sectors by three major leaf veins (Figure 3).</li>
	<li>Tie a small lightweight cardboard tag around the petiole of the leaf containing marked nymph. Number the tag and include notation for the plot or treatment number depending on the experimental design (Figure 2B).</li>
	<li>Tie a 1 m long length of flagging tape around the main stem near the top of the plant. Use a bow-style tie so that the tape can be easily relocated as necessary to keep it visible for repeated visits to this location in the field.</li>
	<li>Record leaf number and positional information on a portable analog or electronic notebook (Table 1). Positional information uses the sectors of the leaf to more finely note location (e.g., 1-1, 1-2, 1-4 denote nymphs in sectors 1, 2 and 4 on leaf #1).</li>
	<li>To ensure that settled 1st instar nymphs and not crawlers are marked, go back to each marked leaf and observe the marked insect about 1-2 h after the initial set-up. Remarking of settled nymphs may be necessary.</li>
	<li>Establish each cohort on a single day. 
	NOTE: A cohort in a given plot is comprised of a minimum of 50 nymphs total. No more than one leaf is used per plant to better distribute the members of the cohort. Depending on insect density, this may comprise anywhere from 13 leaves with 3-4 nymphs each or 50 leaves with one nymph each. The individual plants are selected to distribute the marked leaves along as much of the row as possible. Because of the 1st instar crawler stage, the eggs marked in protocol 2 are not the same insects that are then followed as nymphs. Thus, no crawler mortality is measured and the life table is slightly disjointed in time because egg and nymph cohorts are typically established on the same day. Research has shown the crawler mortality is negligible and can be essentially ignored30.</li>
</ol>
4. Observation and Recording of Egg Hatch and Mortality
<ol>
	<li>After 8-10 d (28-32 &#176;C; average Arizona summer conditions) after establishment of egg cohorts, collect leaves containing marked eggs and return to the laboratory for observation under a dissecting microscope. Eggs are too small to clearly evaluate mortality and causes of mortality in the field.</li>
	<li>Determine causes of death for eggs and record in the notebook initiated at cohort establishment (Table 1). 
	NOTE: Death is characterized as dislodgement, predation or inviability. Dislodgement: the egg is missing due to weather events (wind, blowing dust, rain) or chewing predation. Predation: sucking predators leave behind a collapsed chorion (Figure 4K). Hatched eggs can appear collapsed, but there will be a vertical slit in the egg chorion. Use a minuten pin to tease the chorion on the leaf under the microscope to look for this slit. Inviability: eggs fail to hatch after the 8-10 d period and are a dark tan color. Under Arizona summer conditions (28-32 &#176;C) eggs would normally hatch in 5-7 days. This may differ in other regions and adjustments may be necessary in collection time from the field.</li>
</ol>
5. Observation and Recording of Nymphal Development and Mortality
<ol>
	<li>One to two days after cohort establishment, use a 15X lens to assess the development of nymphs and to assign a cause of mortality if dead. Make observations at least three times per week (every other day).
	<ol>
		<li>Use relative size (Figure 1C-G) and time after establishment to assess instar. 
		NOTE: There are four nymphal instars and development is rapid under Arizona summer conditions (28-32 &#176;C) with each of the first three stages lasting around 2 d and the final stage lasting 3-5 d (total nymphal development 2 wk or less). New observers should learn instar sizes by observing insects reared in the laboratory or greenhouse on the host plant of interest. The relative volume in the abdomen of the bacteriosomes (symbiont harboring organs of the whitefly) relative to overall body size is a helpful indicator of nymphal instar (Figure 1C-G). Newly molted nymphs are very flat and translucent. Nymphs ready to molt are more turgid, domed in profile, and opaque in appearance.</li>
		<li>Determine causes of death for nymphs and record in the notebook initiated at cohort establishment (Table 1). 
		NOTE: Death is characterized as dislodgement, parasitism, predation or unknown depending on instar (Figure 4). Dislodgement: nymphs of any stage are missing due to weather events (wind, blowing dust, rain) or chewing predation. Estimate the stage of dislodged nymphs as the average stage of dead and live nymphs on a given observation date. Parasitism: only observable in 4th instar nymphs. The paired yellowish bacterisomes are displaced by the developing parasitoid larva (Figure 4A); the larval stage is sometimes visible (Figure 4C). The pupal stage of the parasitoid is distinctive and genera specific (Figure 4B, 4D). Predation: sucking predators will evacuate the contents of the nymph and leave behind a collapsed cadaver (Figure 4G-4I). Rarely, a chewing predator may leave evidence (Figure 4J). Unknown: death that cannot be attributed to one of the above causes. In humid environments, fungal disease may be an additional cause of death. This category might also include nymphs killed by parasitoid host-feeding. Nymphs that survive emerge as adults leaving a distinctive t-shaped slot in the exuviae (Figure 1H)</li>
	</ol>
	</li>
	<li>Record development stage (if alive) and cause of death and stage in the notebook initiated at cohort establishment (Table 1).</li>
	<li>Once all the nymphs being observed on a single leaf have either died or emerged as an adult whitefly, collect the leaf and return to the laboratory. Use the higher magnification of a dissecting microscope to confirm that the cause of death noted in the field is accurate and make any corrections. 
	NOTE: Not every non-dislodged dead insect will remain on the leaf over the typical two-week observation period and so some verifications will not be possible.</li>
</ol>
6. Data Summary and Analyses
<ol>
	<li>Consult resources available to help in constructing life tables from the data collected2,8,11,31. An example life table is presented as Table 2. 
	NOTE: Robust life table analyses require multiple independent life tables conducted over time and/or different sites. For a multivoltine insect like B. tabaci this could be multiple life tables over the course of a single season and/or multiple seasons and sites.
	<ol>
		<li>Estimate real mortality (dx/l0) based on the number of insects established at the beginning of the generation. 
		Real mortality=(dx/l0) 
		Where dx is the number dying during stage x and l0 is the number of insects at the beginning of the generation. These mortality rates are additive and the sum of dx over stages estimates the total mortality rate for the generation (Table 2).</li>
		<li>Estimate apparent mortality within a stage (qx) based on the number of insect alive at the beginning of a specific stage (Table 2). Estimate stage-specific qx or factor within stage-specific qx. These rates are additive only within a stage.</li>
		<li>Determine marginal mortality using the formula: 
		MB = dB/(1-dA) 
		Where MB is the marginal rate of mortality factor B, dB is the apparent rate of mortality from factor B and dA is the apparent rate of mortality summed for all mortality factors that can outcompete factor B13,32 (Table 3). 
		NOTE: For sessile insects like whiteflies and many other insects, the multiple causes of death within a particular life stage are not sequential. Instead they act contemporaneously and so estimation of marginal rates is required to accurately estimate stage-specific rates of mortality from any one cause32,34. For example, a parasitoid may attack a whitefly nymph. The parasitoid egg might hatch and the larvae may develop in the host. This activity, initially asymptomatic to the observer, does, or would most likely, kill the host insect and should be credited as the cause of death. But in some cases, a predator may attack this same nymph or the nymph may be dislodged from the leaf leading the observer to note the cause of death as predation or dislodgement. Marginal mortality corrects for this.
		<ol>
			<li>Convert marginal stage-specific rates to k-values35 as k = -ln(1-M), where ln is the natural logarithm and M is the marginal mortality rate of interest. k-values are additive and this simplifies further analyses. k-values can be back-converted to proportional mortality rates by 1-e[-k].</li>
		</ol>
		</li>
		<li>Estimate irreplaceable mortality as [1-e(-TotalK)] - [1-e(-{TotalK - kvaluei})]. 
		NOTE: This gives the portion of total generational mortality that would not be realized if a particular mortality factor was removed. For example, how much generational mortality might be lost if predation or parasitism were removed because of an insecticide spray? Irreplaceable mortality estimated in this way assumes that there is no density-dependence in mortality.</li>
		<li>Key factors
		<ol>
			<li>Use a simple graphical analysis to plot the k-value for any one stage, or any one mortality factor (or one mortality factor within one stage) against the Total-K value for the entire generation (Total K = sum of all individual k-values). 
			NOTE: The mortality factor that most closely mimics the pattern of Total-K the best is the key factor, the factor that contribute the most to changes in generational mortality35. A more quantitative method regresses individual k-values on Total-K and identifies the key factor as the one with the greatest slope value3.</li>
		</ol>
		</li>
		<li>Test density dependence by regressing k-values for factors of interest on the natural log of insect population density measured independently (e.g.13). A significant positive slope suggests direct density dependence and a negative slope inverse dependence. 
		NOTE: With additional life table information on adult survival and reproduction, many additional parameters (e.g., generation time, net reproductive rate, instantaneous rates of increase, life expectancy at a given stage, etc.) and analyses (matrix models and elasticity analyses36,37 can be conducted.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

Typically, the development of life tables for multivoltine insects with broadly overlapping generations are constrained to a vertical approach where a population is sampled repeatedly over time and various graphical and mathematical techniques are then used to estimate recruitment to the various stages and infer rates of mortality from changing densities of the various life stages2. The strength of the approach here is that it navigates this limitation by isolating a group of immobile equal-aged i...

Life tables are a common tool with a long history in ecology1,2. Life tables are essentially a schedule of the births and deaths in a population over time and such data can be used to quantify a number of parameters important to understanding and predicting population dynamics. Life tables may also provide information on causes of death that are important to understanding trophic interactions and in developing control strategies for managing pests in agricultural and natural systems. Numerous field-based life tables have been constructed for insects3,4,5, and analyses have provided important insights into the dynamics, regulation and prediction of insect populations in many managed and natural systems6,7,8,9,10,11,12,13,14. The term life table is also often used to describe laboratory based studies that largely examine schedules of births and deaths but under artificial conditions that do not expose the insect to natural mortality forces and realistic environmental variables. Generally, the goal of laboratory studies is to estimate the comparative biotic potential of a species. The focus of the methods described here is for field based investigations that define realized potential relative to the environment.
Life tables can be characterized as horizontal, in which a real cohort of equal aged individuals are followed from the beginning of their lives until death, or vertical, where frequent samples are taken through time of a population with an assumed stable age structure and then vital rates are inferred from mathematically constructed cohorts2,15. The type of life table that can be deployed depends on the nature of the insect. Horizontal life tables can often be developed for univoltine (one generation per year) insects, while such an approach can be very challenging for a multivoltine insect with multiple and widely overlapping generations each year. A host of analytical methods have been proposed and used to develop vertical life tables for insect populations (see Southwood2 for examples). The methodology demonstrated here allows for the development of cohort-based, horizontal life tables in the field for multivoltine insects with specific life history characteristics, notably, the presence of sessile life stages. The method is demonstrated for a key pest in cotton as a model system.
The whitefly, Bemisia tabaci biotype B (= Bemisia argentifolii, Middle East-Asia Minor 116) is a global pest of agriculture that negatively impacts yield and quality in many agronomic and horticultural crops, including protected agricultural systems in temperate regions17. Impacts occur due to phloem feeding that disrupts nutrient flow, disorders of unknown etiology caused by nymphal feeding, transmission of numerous plant viruses and crop quality effects due to the deposition of honeydew18,19. The insect has a broad host range and is multivoltine, having as many as 12-13 generations per year depending on region and available food resources20. Management challenges also are exacerbated by its high reproductive potential, its ability to disperse and migrate within and between agricultural systems, its lack of a quiescent stage (diapause or estivation) and its disposition to rapidly develop resistance to insecticides used for suppression21,22.
Considerable progress has been made in developing integrated pest management (IPM) strategies to effectively and economically manage populations of this pest in affected crops23,24,25. These management systems were predicated on a sound fundamental understanding of the population dynamics of B. tabaci and life tables have been a key technique that have enabled this understanding. In Arizona, life tables have allowed the estimation and identification of important mortality forces for B. tabaci in multiple crop systems13,26, have enabled the measurement of mortality dynamics relative to management strategies including non-target effects of insecticides14, have provided a means of estimating potential functional non-target effects of transgenic cotton producing insecticidal proteins27, have supported rigorous assessment of a classical biological control program28 (Naranjo, unpublished data) and helped to explore the comparative effects of top-down and bottom-up effects on pest dynamics29. All of these applications have deployed the methodology described here. The approach could be useful for the study of insect population ecology in a number of natural and managed systems.

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			<td>Flagging tape</td>
			<td>Gempler, Janesville, Wisconsin USA</td>
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			<td>American Tag Company, Pico Rivera, California USA</td>
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			<td>Ultra fine point marker</td>
			<td>Sanford, Bellwood, Illinois, USA</td>
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			<td>Adorama, New York, NY USA</td>
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			<td>Adorama, New York, NY USA</td>
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</table>

methodology for developing life tables for sessile insects in the field using the whitefly, bemisia tabaci, in cotton as a model system

Life tables provide a means of measuring the schedules of birth and death from populations over time. They also can be used to quantify the sources and rates of mortality in populations, which has a variety of applications in ecology, including agricultural ecosystems. Horizontal, or cohort-based, life tables provide for the most direct and accurate method of quantifying vital population rates because they follow a group of individuals in a population from birth to death. Here, protocols are presented for conducting and analyzing cohort-based life tables in the field that takes advantage of the sessile nature of the immature life stages of a global insect pest, Bemisia tabaci. Individual insects are located on the underside of cotton leaves and are marked by drawing a small circle around the insect with a non-toxic pen. This insect can then be observed repeatedly over time with the aid of hand lenses to measure development from one stage to the next and to identify stage-specific causes of death associated with natural and introduced mortality forces. Analyses explain how to correctly measure multiple mortality forces that act contemporaneously within each stage and how to use such data to provide meaningful population dynamic metrics. The method does not directly account for adult survival and reproduction, which limits inference to dynamics of immature stages. An example is presented that focused on measuring the impact of bottom-up (plant quality) and top-down (natural enemies) effects on the mortality dynamics of B. tabaci in the cotton system.

An example cohort is presented in Table 2 to show a typical presentation and calculation of life table results. The most useful data is captured in the marginal mortality rates for each factor within each stage. By converting these rates to k-values (protocol section 6), stage-specific mortality over all factors and factor-specific mortality over all stages can be easily estimated, as can total generational mortality. This also facilitates irreplaceable mortality, key-fac...

Watch this Scientific Journal Video about Methodology for Developing Life Tables for Sessile Insects in the Field Using the Whitefly, Bemisia tabaci, in Cotton As a Model System at JoVE.com

Methodology for Developing Life Tables for Sessile Insects in the Field Using the Whitefly, Bemisia tabaci, in Cotton As a Model System

Life tables allow quantification of the sources and rates of mortality in insect populations and contribute to understanding, predicting and manipulating population dynamics in agroecosystems. Methods for conducting and analyzing cohort-based life tables in the field for an insect with sessile immature life stages are presented.

Methodology for Developing Life Tables for Sessile Insects in the Field Using the Whitefly, Bemisia tabaci, in Cotton As a Model System

Life tables provide a means of measuring the schedules of birth and death from populations over time. They also can be used to quantify the sources and ...

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11.7K 조회수.  USDA-ARS, Arid-Land Agricultural Research Center. Life tables allow quantification of the sources and rates of mortality in insect populations and contribute to understanding, predicting and manipulating population dynamics in agroecosystems. Methods for conducting and analyzing cohort-based life tables in the field for an insect with sessile immature life stages are presented.

비디오: Methodology for Developing Life Tables for Sessile Insects in the Field Using the Whitefly, Bemisia tabaci, in Cotton As a Model System

Methods for Performing Crosses in Setaria viridis, a New Model System for the Grasses

Setaria viridis is an emerging model system for C4 grasses. It is closely related to the bioenergy feed stock switchgrass and the grain crop foxtail millet. Recently, the 510 Mb genome of foxtail millet, S. italica, has been sequenced 1,2 and a 25x coverage genome sequence of the weedy relative S. viridis is in progress. S. viridis has a number of characteristics that make it a potentially excellent model genetic system including a rapid generation time, small stature, simple growth requirements, prolific seed production 3 and developed systems for both transient and stable transformation 4. However, the genetics of S. viridis is largely unexplored, in part, due to the lack of detailed methods for performing crosses. To date, no standard protocol has been adopted that will permit rapid production of seeds from controlled crosses.
The protocol presented here is optimized for performing genetic crosses in S. viridis, accession A10.1. We have employed a simple heat treatment with warm water for emasculation after pruning the panicle to retain 20-30 florets and labeling of flowers to eliminate seeds resulting from newly developed flowers after emasculation. After testing a series of heat treatments at permissive temperatures and varying the duration of dipping, we have established an optimum temperature and time range of 48 &deg;C for 3-6 min. By using this method, a minimum of 15 crosses can be performed by a single worker per day and an average of 3-5 outcross progeny per panicle can be recovered. Therefore, an average of 45-75 outcross progeny can be produced by one person in a single day. Broad implementation of this technique will facilitate the development of recombinant inbred line populations of S. viridis X S. viridis or S. viridis X S. italica, mapping mutations through bulk segregant analysis and creating higher order mutants for genetic analysis.

Methods for Performing Crosses in Setaria viridis, a New Model System for the Grasses

We have developed a methodology for performing crosses in Setaria viridis (S. viridis). The method involves pruning the panicle prior to a hot water treatment to kill viable pollen. Crosses are performed following a well-controlled growth regime and typically result in the recovery of 1 to 7 cross-pollinated seed/s per panicle.

에 십자가를 수행하는 방법<em&gt; 강아지풀</em목초에 대한&gt;, 새로운 모델 시스템

Setaria viridis is an emerging model system for C4 grasses. It is closely related to the bioenergy feed stock switchgrass and the grain crop foxtail ...

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A Flow-through Exposure System for Evaluating Suspended Sediments Effects on Aquatic Life

This paper describes the Fish Larvae and Egg Exposure System (FLEES). The flow-through exposure system is used to investigate the effects of suspended sediment on various aquatic species and life stages in the laboratory by using pumps and automating delivery of sediment and water to simulate suspension of sediment. FLEES data are used to develop exposure-response curves between the effects on aquatic organisms and suspended sediment concentrations at the desired exposure duration. The effects data are used to evaluate management practices used to reduce the interactions between aquatic organisms and anthropogenic causes of suspended sediments. The FLEES is capable of generating total suspended solids (TSS) concentrations as low as 30 to as high as 800 mg/L, making this system an ideal choice for evaluating the effects of TSS resulting from many activities including simulating low ambient levels of TSS to evaluating sources of suspended sediments from dredging operations, vessel traffic, freshets, and storms.

A robust and flexible flow-through exposure system designed to maintain sediment in suspension is presented. The system is used to investigate the effects of suspended sediment on various aquatic species and life stages in the laboratory.

수산 생명에 일시 중지를 평가하면 퇴적물 효과에 대한 플로우를 통해 노출 시스템

This paper describes the Fish Larvae and Egg Exposure System (FLEES). The flow-through exposure system is used to investigate the effects of suspended ...

Controlled-release of Chlorine Dioxide in a Perforated Packaging System to Extend the Storage Life and Improve the Safety of Grape Tomatoes

A controlled-release chlorine dioxide (ClO2) pouch was developed by sealing a slurry form of ClO2 into semipermeable polymer film; the release properties of the pouch were monitored in containers with or without fruit. The pouch was affixed to the inside of a perforated clamshell containing grape tomatoes, and the effect on microbial population, firmness, and weight loss was evaluated during a 14 day storage period at 20 &#176;C. Within 3 days, the ClO2 concentration in the clamshells reached 3.5 ppm and remained constant until day 10. Thereafter, it decreased to 2 ppm by day 14. The ClO2 pouch exhibited strong antimicrobial activity, reducing Escherichia coli populations by 3.08 log CFU/g and Alternaria alternata populations by 2.85 log CFU/g after 14 days of storage. The ClO2 treatment also reduced softening and weight loss and extended the overall shelf life of the tomatoes. Our results suggest that ClO2 treatment is useful for extending the shelf life and improving the microbial safety of tomatoes during storage without impairing their quality.

Here, we describe a protocol for the application of a novel, slow-release ClO2 product that reduces spoilage and extends the shelf life of fresh fruit. The slow-release ClO2 product was added to standard commercial grape tomato packaging and tested against Escherichia coli and Alternaria alternata.

천공 포장 시스템에서 이산화 염소의 제어 릴리스는 저장 수명을 연장하고 포도 토마토의 안전을 개선하기 위해

A controlled-release chlorine dioxide (ClO2) pouch was developed by sealing a slurry form of ClO2 into semipermeable polymer film; the release properties ...

Techniques for Investigating the Anatomy of the Ant Visual System

This article outlines a suite of techniques in light microscopy (LM) and electron microscopy (EM) which can be used to study the internal and external eye anatomy of insects. These include traditional histological techniques optimized for work on ant eyes and adapted to work in concert with other techniques such as transmission electron microscopy (TEM) and scanning electron microscopy (SEM). These techniques, although vastly useful, can be difficult for the novice microscopist, so great emphasis has been placed in this article on troubleshooting and optimization for different specimens. We provide information on imaging techniques for the entire specimen (photo-microscopy and SEM) and discuss their advantages and disadvantages. We highlight the technique used in determining lens diameters for the entire eye and discuss new techniques for improvement. Lastly, we discuss techniques involved in preparing samples for LM and TEM, sectioning, staining, and imaging these samples. We discuss the hurdles that one might come across when preparing samples and how best to navigate around them.

This article outlines a suite of techniques in light and electron microscopy to study the internal and external eye anatomy of insects. These include several traditional techniques optimized for work on ant eyes, detailed troubleshooting, and suggestions for optimization for different specimens and regions of interest.

개미 비주얼 시스템의 조사 기법

This article outlines a suite of techniques in light microscopy (LM) and electron microscopy (EM) which can be used to study the internal and external eye ...

Harvesting Venom Toxins from Assassin Bugs and Other Heteropteran Insects

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

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

독 독 소 암살자 버그와 다른 Heteropteran 곤충에서 수확

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

A Precise and Autonomous System for the Detection of Insect Emergence Patterns

Existing systems to measure insect emergence patterns have limitations; they are only partially automated and are limited in the maximum number of emerging insects they can detect. In order to obtain precise measurement of insect emergence, it is necessary for systems to be semi-automated and able to measure large numbers of emerging insects. We addressed these issues by designing and building a system that is automated and can measure emergence of up to 1200 insects. We modified the existing &#34;falling-ball&#34; system using Arduino microcontrollers to automate data collection and expand the sample size through multiple data channels. Multiple data channels enable the user to not only increase their sample size, but also allows for multiple treatments to be run simultaneously in a single experiment. Furthermore, we created an R script to automatically visualize the data as a bubble plot, while also calculating the median day and time of emergence. The current system was designed using 3D printing so the user can modify the system to be adjusted for different species of insects. The goal of this protocol is to investigate important questions in chronobiology and stress physiology, using this precise and automated system to measure insect emergence patterns.

Measurement of insect emergence patterns requires precision. Existing systems are only semi-automated and sample size is limited. We addressed these issues by designing a system using microcontrollers to precisely measure the time of emergence of large numbers of emerging insects.

곤충 출현 패턴의 검출을 위한 정확 하 고 자치 시스템

Existing systems to measure insect emergence patterns have limitations; they are only partially automated and are limited in the maximum number of ...

Visualizing Efficacy of Pesticides Against Disease Vector Mosquitoes in the Field

Efficacy of public health pesticides targeting nuisance and disease-vector insects such as mosquitoes, sand flies, and filth-breeding flies is not uniform across ecological zones. To best protect public and veterinary health from these insects, the environmental limitations of pesticides need to be investigated to inform effective use of the most appropriate pesticide formulations and techniques. We have developed a research program to evaluate combinations of pesticides, pesticide application equipment, and application techniques in hot-arid desert, hot-humid tropical, warm and cool temperate, and urban locations to derive pesticide use guidelines specific to target insect and environment. To these ends we designed a system of protocols to support efficient, cost-effective, portable, and standardized evaluation of a diverse range of pesticides and equipment across multiple environments. At the core of these protocols is the use of an array of small cages with colony-reared sentinel mosquitoes (adults and immatures) and sand flies (adults), strategically arranged in natural habitats and exposed to pesticide spray. Spatial and temporal patterns of pesticide efficacy are derived from percent mortality in sentinel cages, then mapped and visualized in a geographic information system. Maps of sentinel mortality data may be statistically compared to evaluate relative efficacy of a pesticide across multiple environments, or to study multiple pesticides in a single environment. Protocols may be modified to accommodate a variety of scenarios, including, for example, the vertical orientation of sentinels in canopy habitats or simultaneous testing of ground and aerial application methods.

The efficacy of public health pesticides targeting nuisance and disease-vector insects is not uniform across different ecological zones. Here we present a system of techniques using captive vector insects as sentinels for pesticide efficacy to derive electronic maps supporting the standard evaluation of pesticides across multiple environments.

시각화 분야에서 질병 벡터 모기에 대 한 살충제의 효능

Efficacy of public health pesticides targeting nuisance and disease-vector insects such as mosquitoes, sand flies, and filth-breeding flies is not uniform ...

A Loop-mediated Isothermal Amplification (LAMP) Assay for Rapid Identification of Bemisia tabaci

The whitefly Bemisia tabaci (Gennadius) is an invasive pest of considerable importance, affecting the production of vegetable and ornamental crops in many countries around the world. Severe yield losses are caused by direct feeding, and even more importantly, also by the transmission of more than 100 harmful plant pathogenic viruses. As for other invasive pests, increased international trade facilitates the dispersal of B. tabaci to areas beyond its native range. Inspections of plant import products at points of entry such as seaports and airports are, therefore, seen as an important prevention measure. However, this last line of defense against pest invasions is only effective if rapid identification methods for suspicious insect specimens are readily available. Because the morphological differentiation between the regulated B. tabaci and close relatives without quarantine status is difficult for non-taxonomists, a rapid molecular identification assay based on the loop-mediated isothermal amplification (LAMP) technology has been developed. This publication reports the detailed protocol of the novel assay describing rapid DNA extraction, set-up of the LAMP reaction, as well as interpretation of its read-out, which allows identifying B. tabaci specimens within one hour. Compared to existing protocols for the detection of specific B. tabaci biotypes, the developed method targets the whole B. tabaci species complex in one assay. Moreover the assay is designed to be applied on-site by plant health inspectors with minimal laboratory training directly at points of entry. Thorough validation performed under laboratory and on-site conditions demonstrates that the reported LAMP assay is a rapid and reliable identification tool, improving the management of B. tabaci.

A Loop-mediated Isothermal Amplification LAMP Assay for Rapid Identification of Bemisia tabaci

This paper reports the protocol for a rapid identification assay for Bemisia tabaci based on loop-mediated isothermal amplification (LAMP) technology. The protocol requires minimal laboratory training and can, therefore, be implemented on-site at points of entry for plant imports such as seaports and airports.

루프-중재 등온 증폭 (램프) 분석 결과 Bemisia tabaci 의 빠른 식별

The whitefly Bemisia tabaci (Gennadius) is an invasive pest of considerable importance, affecting the production of vegetable and ornamental crops in many ...

An Induction System for Clustered Stomata by Sugar Solution Immersion Treatment in Arabidopsis thaliana Seedlings

Stomatal movement mediates plant gas exchange, which is essential for photosynthesis and transpiration. Stomatal opening and closing are accomplished by a significant increase and decrease in guard cell volume, respectively. Because shuttle transport of ions and water occurs between guard cells and larger neighboring epidermal cells during stomatal movement, the spaced distribution of plant stomata is considered an optimal distribution for stomatal movement. Experimental systems for perturbing the spaced pattern of stomata are useful to examine the spacing pattern's significance. Several key genes associated with the spaced stomatal distribution have been identified, and clustered stomata can be experimentally induced by altering these genes. Alternatively, clustered stomata can be also induced by exogenous treatments without genetic modification. In this article, we describe a simple induction system for clustered stomata in Arabidopsis thaliana seedlings by immersion treatment with a sucrose-containing medium solution. Our method is easy and directly applicable to transgenic or mutant lines. Larger chloroplasts are presented as a cell biological hallmark of sucrose-induced clustered guard cells. In addition, a representative confocal microscopic image of cortical microtubules is shown as an example of intracellular observation of clustered guard cells. The radial orientation of cortical microtubules is maintained in clustered guard cells as in spaced guard cells in control conditions.

An Induction System for Clustered Stomata by Sugar Solution Immersion Treatment in Arabidopsis thaliana Seedlings

The goal of this protocol is to demonstrate how to induce clustered stomata in cotyledons of Arabidopsis thaliana seedlings by immersion treatment with a sugar-containing medium solution and how to observe intracellular structures such as chloroplasts and microtubules in the clustered guard cells using confocal laser microscopy.

클러스터 Stomata 애기 thaliana 묘에 설탕 솔루션 집중 치료에 의해 위한 유도 시스템

Stomatal movement mediates plant gas exchange, which is essential for photosynthesis and transpiration. Stomatal opening and closing are accomplished by a ...

A Gnotobiotic System for Studying Microbiome Assembly in the Phyllosphere and in Vegetable Fermentation

The phyllosphere, the above ground portion of the plant that can be colonized by microbes, is a useful model system to identify processes of microbial community assembly. This protocol outlines a system for studying microbial community dynamics in the phyllosphere of Napa cabbage plants. It describes how to grow germ-free plants in test tubes with a calcined clay and nutrient broth substrate. Inoculation of germ-free plants with specific microbial cultures provides opportunities to measure microbial growth and community dynamics in the phyllosphere. Through the use of sterile vegetable extract produced from cabbages shifts in microbial communities that occur during fermentation can also be assessed. This system is relatively simple and inexpensive to set up in the lab and can be used to address key ecological questions in microbial community assembly. It also provides opportunities to understand how phyllosphere community composition can impact the microbial diversity and quality of vegetable fermentations. This approach for developing gnotobiotic cabbage phyllosphere communities could be applied to other wild and agricultural plant species.

A method of growing germ-free Napa cabbages has been developed which enables researchers to evaluate how single microbial species or multispecies microbial communities interact on cabbage leaf surfaces. A sterile vegetable extract is also presented which can be used to measure shifts in community composition during vegetable fermentation.

필로스피어와 채소 발효에서 미생물군유전체 조립을 연구하기 위한 노토바이오틱 시스템

The phyllosphere, the above ground portion of the plant that can be colonized by microbes, is a useful model system to identify processes of microbial ...