Bureau of Reclamation Banner

Technical Service Center
Environmental Applications and Research Group — Publications

Saltcedar Biocontrol Activities at the Bureau of Reclamation Technical Service Center

Technical Memorandum No. 8220-99-01
by
Debra Eberts

1999

Saltcedar Biocontrol Activities
Chronology and Description of Activities
Methods and Observations
Future Plans
Acknowledgments


Saltcedar Biocontrol Activities

Saltcedar (Tamarix sp., especially ramosissima) is a nonnative shrubby tree (photo 1) that has become a noxious weed in riparian systems of the western United States. Native vegetation such as cottonwoods and willows is replaced by dense monocultures of saltcedar. This tree depletes groundwater resources, and increases soil salinity, soil erosion, and flooding events. Where this plant has invaded Bureau of Reclamation systems, it creates an expensive management challenge to effectively manage water and habitat resources.

Traditional chemical and mechanical methods have been successful in managing saltcedar at some sites, but these methods are expensive and labor-intensive. Dr. C. Jack DeLoach of the U.S. Department of Agriculture, Agricultural Research Service (USDA-ARS) has been researching biocontrol insects for saltcedar at the Grassland, Soil, and Water Research Laboratory in Temple, Texas. He met with Reclamation in 1996 to evaluate cooperative research and release sites on Reclamation property near Pueblo, Colorado. However, field releases of insects are on hold until an Environmental Assessment is returned by USDA-APHIS (U.S. Department of Agriculture, Animal and Plant Health Inspection Service) with a finding of no significant impact (FONSI). This requirement became necessary when the southwestern subspecies of the willow flycatcher, which has been found to nest in saltcedar in some areas, was designated as an endangered species.

In 1997, Dr. DeLoach and Reclamation received a permit from USDA-APHIS-PPQ to begin a small caged field study at the Pueblo, CO site if certain security measures were taken.

Chronology and Description of Activities

May-June, 1997 Three experimental sites were prepared. The overall site was chosen based on its ease of access to researchers, and improbability of disturbance by others. It also has the dry soils required for the insects to complete their life cycle, with very little possibility of flooding. Clumps of saltcedar bushes were chosen based on the feasibility of fitting into a tent with a 12 foot by 12 foot footprint. We chose clumps consisting of several individual trees, so that the death of one would have less impact. These trees were trimmed to a height of about 5 feet (photo 2).

Trenches were dug to fit the support framing (photos 3, 4), and two 12 ft x12 ft x 6 ft Lumite Saran® 20x20 mesh tents were used on each assembly to give double-walled tents. The bottom flaps were sandwiched between wooden boards and buried in the ground to a depth of about 12 inches by refilling the trenches. Each tent was enclosed by a security chainlink fence with a locking gate (photo 5).

June 24/25, 1997 We received a shipment of 108 eggs of Diorhabda elongata elongata (Kazakhstan, Chilik) from Dr. DeLoach. These eggs had been laid over the period June 18 to 23 at Dr. DeLoach's laboratory in Texas. On June 25, the eggs were taken to the study site, attached to suitable branches in tent #2 (site photos), and covered with mesh bags to confine the larvae and make their relocation and observation easier (photo 7). Small roofs were made from reflective insulation to shelter the bags and decrease mortality caused by wet bags after a rain (photo 8).

June 30, 1997 The first larva was found. The weather had been very hot (about 100°F) and dry since the eggs were put out.

July 8, 1997 Survey of all mesh bags showed there were 70 larvae (65% yield from eggs), mostly first instars. Removing and retying the mesh bags to check and count larvae and pupae did cause a few casualties each week. Weather is still hot (about 95°F) and dry.

July 16, 1997 We decided to remove bags permanently from about half the larvae to allow them to pupate where they chose. This left 30 larvae still enclosed in mesh bags. By this time, most larvae were second or third instar. Temperatures were about 100-105°F.

July 23, 1997 Noticed prepupae ("C"-shaped, inactive larvae). Temperatures were about 100°F.

July 29, 1997 Did not remove remaining bags to do evaluations; saw pupae within. Heavy rains and cooler temperatures (80s) since the weekend.

August 5, 1997 Checked all remaining bags, found 12 adults and 10 pupae. Put each group into a mesh bag over new foliage. Heavy rain last night, highs today low 70s.

August 12, 1997 The pupae still enclosed in mesh bags developed into 3 live adults. Seven of the pupae that had been moved to new bags August 5 became deformed and dead adults.

August 19, 1997 No eggs had been laid after several weeks, so decided to release these adults from the bags into the tent.

October 22, 1997 One adult beetle was seen on the saltcedar foliage.

October 24, 1997 In the past, late spring snowstorms have caused collapsed tents in our work with other biocontrol insects. Therefore, we constructed metal carports to prevent snow from ripping or collapsing the tents in Pueblo. These carports were received and erected in early to mid-October (photo 6). Steve Williams and Roy Vaughan from the Pueblo Field Office erected the carports before snow arrived. Over October 24-26, several feet of snow fell, accompanied by high winds and near-zero temperatures. The tents remained secure.

Late April and May, 1998 Trees were pruned, carports removed and weekly checks were made for Diorhabda adults. No adults were found before May 30, even though foliage was abundant (4 inches long on May 6). It was possible the shade from carports was keeping soil cool and delaying emergence, but removal of the carports did not result in appearance of insects in tent #2.

May 29/30, 1998 A shipment of eggs was received from Dr. DeLoach. These eggs were laid over the period May 19 to May 24. Approximately 1,000 eggs were shipped, but the package was damaged in shipment and we counted only 585 undamaged eggs. These were glued to Nalgene® paper using Elmer's Glue® and released into mesh bags in tent #1. Papers containing a total of about 25 eggs were placed into mesh bags on May 30. In 1997, the pieces of paper had been glued onto branches, but this technique was changed to carefully placing them into the bag after it was placed over the foliage.

Eggs received this season were of Diorhabda elongata deserticola Chen, Order Coleoptera: Family Chrysomelidae. Determined by Li Bao-ping, Xinjiang Agric. Univ (XAU). Collected in China: Xinjiang Prov., Fukang by Li Bao-ping, XAU.

June 2/4, 1998 Another set of about 800 eggs (laid May 27 to June 1) was received June 2 and taken to Pueblo on June 4. The eggs were glued to paper and placed into bags as before. Eggs were beginning to hatch as they were put into mesh bags on the tree in tent #2, and numerous first instar larvae were present. The release rate was about 35-40 eggs/bag.

June 22, 1998 Monitoring of a sample of the bags showed that tent #1 had about a 28% hatch rate, and tent #2 was about 16%. Larvae in both tents were second or third instar. On this date, leafhoppers were also beginning to hatch inside and outside the bags and damage foliage. We put a fresh set of yellow sticky cards into tent #3 to catch leafhoppers.

July 7, 1998 Found a few adults in bags, many pupae. In a few bags, foliage was significantly damaged by leafhoppers so fresh foliage was added. Temperatures have been near 100°F.

July 15, 1998 Temperatures again near 100°F. Foliage in bags in tent #1 entirely dead. Foliage in bags in tent #2 about 90% dead. Many dead and live leafhoppers inside bags. Generally, foliage outside of bags appears more healthy than inside the bags (this probably due more to leafhoppers than Diorhabda) (photo 12). Collected adults and took all adults and pupae to tent #1. Set up one bag with all pupae and 2 bags with 12 adults each, then released the rest of the adults into tent #1. The pupae were transferred without touching; they were "poured" from their original mesh bags into the new bag.

July 28, 1998 Visited site with Juli Gould, USDA-APHIS-PPQ, Phoenix, AZ. No insects or signs of insects were seen in tent #2. Scattered ˝ bale of straw in each of tents #1 & #2 to provide more insulation for overwintering insects. Found adults in tent #1 laying eggs and some larvae hatched from these eggs (mostly first instar, a couple second instar larvae). Found about 150 unhatched eggs plus larvae without searching thoroughly. Of the bags containing adults and pupae: eggs were found in one bag of adults and the bag which contained pupae. No deformed or dead adults or pupae were found as a result of transferring pupae.

August 4, 1998 Divided another bale of straw between tents #1 and #2. Still no sign of insects in tent #2, so transferred some from tent #1: collected 2 males and 4 females (mating in collection vial), and eight branch cuttings with eggs (estimate about 100 eggs total). Adults were released into the tent and cuttings were not bagged, just attached with clothespins.

September 9, 1998 Tent #1 had numerous adults; observed one third instar larva. Only about 5% of tree foliage destroyed by leafhoppers. Destroyed about 8 jumping spider nests (Phidippus sp.) and removed the spiders. In tent #2 about 90-95% of the foliage had been killed, probably by leafhoppers. Combined with an explosion of too many Phidippus to begin to count, chances for survival of larvae did not look good. Tent #3 had abundant leafhoppers, but only 5-10% of foliage damaged.

October 5, 1998 Carports put back over tents.

October 21, 1998 Tent #1: foliage good, no Diorhabda seen. Tent #2: regrowth of foliage, no Diorhabda seen. Tent #3: Some green foliage, but much current leafhopper damage. In site area outside of tents about 90% of saltcedar turned yellow in the past week due to temperature (has been down to 26° a few times).

Methods and Observations

Egg/Larval handling
Eggs obtained from source cultures of Diorhabda should be collected and shipped with foliage or mesh bag attached to avoid damaging the eggs. They should also be shipped overnight and securely padded, in cardboard boxes rather than padded envelopes.

Eggs should be placed at field sites as soon as possible after receipt, preferably no later than the next day. We used Elmer's Glue® and toothpicks to glue the eggs to small (1 inch x ˝ inch) cards cut from Nalgene® paper (photo 10). We used as little glue as possible and tried to avoid contact with the eggs. We usually glued the foliage to the card, but if the eggs had fallen off the foliage, the glue was allowed to become tacky before placing the eggs; otherwise they tended to "sink". The egg count was written on each card so that we could limit the number of eggs per mesh bag to 20-25. Egg cards were taped to a sheet of paper and transported to the field site in a multi-drawer "small parts" organizer with drawers only slightly larger than the sheet of paper.

The first step at the field site was to make the number of foliage bundles we needed. A number of stems would be gathered together and twist-tied at least a foot from the tips. Once these were all prepared then we would place the mesh bag over the foliage and carefully insert the eggs. Eggs could easily be damaged at this point. Care must be taken to ensure that the eggs maintain the paper side toward the bag and do not scrape on any twigs. The mouth of the bag was tied as tightly as possible with another twist-tie. A Nalgene® label can be attached to this twist-tie if you want to keep records on egg numbers or date.

After this point, it would be best to handle the insects as little as possible. If you do want to examine the insects up close, a large white cafeteria-style tray should be held under the bag as you open and remove the bag and search through the foliage. This is difficult for one person to do alone, so working in pairs is recommended. The larvae do fall off, but they quickly climb back onto foliage offered to them. An aspirator should be handy to collect any adults.

We did not note any mortality due to pupation inside the mesh bags as long as pupae were not manipulated. Frequently, larvae would crawl up to the tightest part of the bag to pupate (where it was tied), but pupae were also visible just laying in a corner of the bag, not heavily covered by foliage. If the insects will be pupating inside the tent but not in bags, caution should be used when walking around inside the tent when there might be pupae in the soil and litter.

Yield from eggs
In the 1997 shipment, 108 eggs gave 70 larvae (65% hatch). From larvae retained in the bags (30) we obtained 14 healthy adults plus 7 adults which were deformed due to handling. This would give an estimated 45% yield from egg to adult. Methods of egg harvest and handling of eggs and larvae in 1997 did not cause unacceptable levels of mortality, although some mortality of larvae or pupae was caused by handling and the removal/replacement of bags for observations.

In 1998 the yields were much lower. Only 181 adults were obtained from 1385 eggs, a 13% yield. It is likely that most of the mortality occurred in the egg stage. Techniques used to collect the eggs may have slightly damaged their shells. When the eggs were then treated with zephiran chloride to prevent Beauvaria (fungus) infection, the chemical may have killed a greater number of eggs than if the shells had been undamaged. Additionally, one set of eggs was hatching as it was being placed in the cages. While we attempted to handle the newly-emerged first instar larvae as gently as possible, there were likely a number of mortalities due to the transfer.

Larval stages
Data from Dr. DeLoach (Effects of Biological Control of Saltcedar (Tamarix ramosissima) on Endangered Species, Biological Assessment, April 1997) on duration of larval stages and size of larvae is excerpted as follows:

Table 1. Size and duration of the immature stages of Diorhabda elongata.*

stage/instar
egg
first
second
third
prepupa ("C"-shape)
pupa
duration (days)
ca. 10
4.92 ± 1.04
4.77 ± 1.04
7.39 ± 1.66
4.78 ± 1.52
7.08 ± 0.76
body length (mm)
--
2
4
9
--
--

*The above data was collected in the quarantine laboratory at Temple, Texas, during the late summer of 1992, at room temperature of 25°±3°, 16:8 day-night photoperiod.

The following table compares DeLoach's laboratory findings of development times to findings at the Pueblo field site. A range in days is shown because eggs were laid over a number of days and pooled together. Further error of up to 6 days as compared to DeLoach values may be added because the site was not visited more than once per week. As can be seen, Pueblo ranges fall within the ranges seen by DeLoach, although the entire cycle from egg to adult appears to average a few days longer. This may be due to lower temperatures caused by geographical location or shading by the tent enclosures.

Table 2. Comparison of laboratory and field values for duration of the
immature stages of Diorhabda elongata.*


stage/instar
egg
first
second
third
prepupa
pupa
adult
Laboratory stage
at day #

0-10
10-15
15-20
20-28
28-33
34-41
42

1997
0-12
12-20
--
--
30-35
36-41
43-55
Field Stage at day #
1998 #1
--
--
--
29-34
--
44-49
44-52

1998 #2
--
--
18-20
--
--
33-35
41-56

*Laboratory data collected by Dr. DeLoach in the quarantine laboratory at Temple, Texas, during the late summer of 1992, at room temperature of 25°±3°, 16:8 day-night photoperiod. Field data from trials inside tents at the Pueblo, Colorado test site.

A second generation of insects resulted in 1998 by day 65-70 when many eggs and a number of first instar larvae were found. These eggs had been laid by insects allowed to move freely about the tent. A percent yield from these eggs was not determined, although larvae were numerous.

Leafhopper competition
Short of repeated insecticidal treatment or fumigation, it is impossible to rid plant cultures of the leafhopper Opsuis stactogalus. This accidentally-introduced leafhopper is specific to saltcedar and can be devastating to foliage inside tents or bags. We have set up multiple tents at the Pueblo field site so that Diorhabda can be transferred to another tent if the foliage in their tent is destroyed by leafhoppers.

We did attempt to reduce leafhopper numbers in the extra tent by using yellow sticky cards placed out just before leafhopper emergence and replaced once during the season. We used about 50 (3 inch x 5 inch) cards in the tent each time. It is not clear whether this actually helped, because foliage damage in this treated tent was visually estimated at 5-10%, while the two tents with Diorhabda and no sticky cards had damage estimated at 5% (tent with most of Diorhabda), and 90-95% (tent with few Diorhabda).

Leafhoppers were first noted in mid-June, significant foliage damage in a few mesh bags was seen 2 weeks later, and foliage in all bags was 90-100% brown by the next week. Dead Diorhabda adults found in bags on July 15 were collected from bags where there had been almost complete destruction of foliage by leafhoppers (photo 12). A handful of dead leafhoppers was poured out of some of these bags. After emergence of leafhoppers, bags should probably be checked more than once a week to keep Diorhabda mortality to a minimum.

Overwintering conditions
In most biocontrol experiments where outdoor tents are used, the tents are taken down after the mobile insect stages have died or gone into a "resting" stage for the winter. Regulations imposed on this field test required that tents remain in place over winter. Previous personal experience with late spring snows causing collapsed tents in other projects led us to anchor a carport over each tent in Pueblo (photo 6) before the first snow.

The carports cause unnatural conditions inside the tents while they are in place. Light and precipitation are both decreased. Lack of precipitation and reduced winds led to the trees retaining almost all of the dead foliage and reduced litter on the ground for the insects. In 1997, these conditions in combination with the presence of no more that 50 adults (that had not produced any eggs) likely led to a failure of any Diorhabda surviving the winter.

For the winter of 1998, it will still be necessary to have the tents and carports in place. Litter was supplemented with straw which will be sprayed with water (but not soaked) several times over the season. Spraying will be done from the door without entering the cage.

Behavior/Monitoring
Monitoring of the behavior of the insects wasn't possible until July 1998 when there was a healthy population of all insect stages free within the tent. Behavior may still be affected by the enclosure, but some observations may remain valid. Obviously, it is much easier to find eggs, larvae, and adults in an enclosure than in a field situation. All stages were easy to see and identify in an enclosure, but the first instar's shed skin may provide a good visual cue for locating Diorhabda in the field. The first instar larvae primarily left their skins attached to small outer twigs of foliage (photo 13). With the light behind them, these skins (or the larvae themselves) are easy to see from a distance. In the absence of rain or wind, they should persist long enough to show up under regular surveys.

When eggs were laid by insects allowed to move freely about the tent (July/August 1998), it was noticed that no eggs or larvae occurred on the north side of the trees in this one trial. Further observations should be made to determine if the insects prefer hotter, more sunny exposures for larval development.

Future plans

In late December 1998, the Assistant Director of the U.S. Fish and Wildlife Service communicated to USDA-APHIS-PPQ that they concur that experimental releases on the 13 selected sites will not adversely affect the southwestern willow flycatcher. There is potential that APHIS will soon be able to approve an Environmental Assessment and give permits for release at the experimental sites.

Work will continue to develop and test plans for monitoring vegetation and insects, and methods for rearing large numbers of insects. Cooperation will continue with USDA-ARS to develop and share these methods.

Acknowledgments

We would like to express our appreciation for the contributions and support from the following:

Partnership Resources
Bureau of Reclamation Research Program WATER Project EE007, "Development of Improved Aquatic Site Pest Management Methods"
Bureau of Reclamation Program Analysis Office

Cooperators
Bureau of Reclamation, Pueblo Field Office, Pueblo, CO
Bureau of Reclamation, Eastern Colorado Area Office, Loveland, CO
Dr. C. Jack DeLoach, USDA-ARS, Grassland, Soil, and Water Research Lab, Temple, TX
Colorado Department of Agriculture, Denver, CO


[Bureau of Reclamation Home Page]