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Methodology for Developing Life Tables for Sessile Insects in the Field Using the Whitefly, Bemisia tabaci, in Cotton As a Model System
Chapters
Summary November 1st, 2017
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.
Transcript
The overall goal of this cohort-based life table approach is to directly quantify the rate of mortality of an insect population in the field and identify the source of mortality. This method can help us answer key questions in population dynamics such as understanding factors contributing to mortality in populations and ultimately the rate at which populations grow and decline. The main advantage of this technique is that it directly measures sources and rates of mortality from multiple causes under realistic conditions in the field.
We first had the idea for this method when we learned the insect is immobile in its immature stages. This allows us to directly follow the fate of individual insects. Demonstrating the procedure will Isadora Bordini, a graduate student, and Francisco Bojorquez, a lab technician.
Prepare 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. 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. Pick a sampling site three to four meters into the row. To simplify later observations, select one row to establish egg cohorts and the second to establish nymph cohorts.
An egg cohort must include at least 50 eggs. As a consequence of behavior and plant growth, the life stages of Bemisia tabaci are generally distributed vertically in the canopy with the eggs near the top and progressively older nymphal stages below. Use an 8x hand lens to search for newly-laid eggs on the underside of leaves.
Ideally, the hand lens should have a hole in the side that the pen can fit through. Find one leaf with eggs for each plant in the study. Next, use a 15x lens to make a more careful examination.
Eggs darken to a tannish color as they mature so fresh eggs will have a bright white coloration and stand out from the older eggs. The most critical aspect of this protocol is to correctly identify the newly-laid eggs and newly-settled first instar nymphs. It is vital that the life table observations start at the very beginning of each life stage for accurate results.
After finding an egg, use a nontoxic ultra fine point black permanent marker to draw a small circle around the egg. Keep the circle small so no other eggs will get laid within it. On the same leaf, mark up to four eggs in the four different sectors.
On a cotton leaf, there are four sectors divided by the major leaf veins. Only select one egg per leaf sector. Identify the marked leaf using a small lightweight cardboard tag around the petiole of the leaf with notation for the plot or treatment number depending on the experimental design.
Then attach a highly visible flag near the top of the plant. Now, move on to finding eggs on the next plant further along the row. Establish the cohort in a single day.
Under normal Arizona summer conditions, the leaves will be ready to collect in eight to 10 days because the eggs hatch in five to seven days. Adjust the time of collection to ensure that all the healthy eggs should have hatched. In the field, eggs are too small to clearly evaluate mortality and causes of death in situ.
Therefore, the causes of mortality like dislodgment, predation, or inviability are determined in the laboratory under a dissection microscope. It is critical to correctly identify the cause of death of the egg or nymph as this affects the ecological inferences that can be drawn about factors affecting population growth and regulation. If the egg is missing due to weather or chewing predation, then it has been dislodged.
If an egg has been rendered into a collapsed chorion, this is classified as predation. Hatched eggs also appear collapsed, but have a vertical slit in the chorion which can be found using a tiny pin. If an egg failed to hatch, then categorize it as inviable.
Use an 8x hand lens to search for newly-settled first instar nymphs on the underside of leaves. Such leaves will be about three to five nodes down from the top of a cotton plant. When any nymphs are found, verify their stage using a 15x lens.
Find at least 50 nymphs for each study. Select plants that are well distributed along the row and establish the cohort in a single day. As before, draw a small circle around the identified nymph.
Mark up to four nymphs per leaf each in a different sector and then proceed to the next plant. Be aware that freshly-hatched first instar nymphs can move several centimeters over before they settle and this can take a few hours. Settled nymphs can be distinguished from crawling nymphs because they are immobile, are more tightly pressed against the leaf, and have a more translucent amber color.
Older first instar nymphs appear swollen and turgid as they near molting. Mark a tag with notation for the plot or treatment number and secure it to the petiole of the leaf. Then attach a highly visible flag near the top of the plant.
Make detailed notes of the position of the nymph including leaf sector information. After an hour or two of identifying nymphs, double check them to ensure that crawling nymphs were not circled. Remark the settled nymphs as needed.
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 or about every other day. Death is characterized as dislodgement, parasitism, predation, or unknown depending on instar.
A typical life table of a cohort provides the marginal mortality rates for each factor within each life stage. By converting these rates to K values, stage-specific mortality over all factors and factor-specific mortality over all of the stages can be easily estimated as can the total generational mortality. Using these techniques in a three-year study, it was found that marginal rates of predation by sucking predators declined significantly when broad spectrum insecticides were applied.
Patterns of irreplaceable mortality were similar. Normally, predation supplied the highest level of irreplaceable mortality, but this declined significantly with insecticide use. Parasitism also supplied low levels of irreplaceable mortality.
Key factor analysis made from the data on four replicate plots were combined into a total of nine cohorts over the three-year study. Ultimately, predation was most closely associated with changes in generational mortality. Once mastered, this technique can be done in about 30 minutes per plot, but this can vary greatly depending on insect density.
While attempting this procedure, it's important to remember to minimize stress on the leaf being manipulated so that plant quality is not altered. Also, a single observer should be used per replicate to block for observer effects and all observers should meet frequently to enhance uniformity of observational criteria. After watching this video, you should have a good understanding of how to set up a life table for an immobile insect in the field including locating, marking, and observing insects to determine causes of death.
Don't forget that working in the field for long periods especially during summer months can be difficult and that proper attire and hydration are necessary to perform the work safely.
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