In December, I updated you on how perennial wildflowers and grasses were establishing in our beneficial insect habitat plots during the 2019 growing season. As I wrote that post, I quickly realized that there was too much good information for just one post. So here’s the rest of the story when it comes to plant establishment – time, money, and weeds.
Before I get started, below is a quick reminder of what our treatments were. You can read all the details here.
Treatment
Description
A
Spring transplant, no mulch
B
Spring transplant with mulch
C
Spring direct seed
D
Buckwheat cover crop, then fall transplant
E – control
Whatever was growing there, just keep it mowed
F
Soil solarization, then fall direct seed
G
Herbicide and tillage, then fall direct seed
Weed control
One thing that has surprised me about this project (although others certainly gave me fair warning) was how big a role weed management plays in establishing habitat for beneficial insects. It’s definitely still a struggle in our plots.
Bryan Brown did weed assessments for us in May and September of 2019. The graph below shows the average percent of the area of each plot covered by either weeds (orange) or beneficial habitat flowers and grasses (blue).
Mulching provided the best weed management when plots were assessed in May of 2019. Each bar shows the average of four plots per treatment, and has an error bar showing variation among these plots (one standard error above or below the average for the treatment).
This was before we did any hand-weeding. By far, the plots that were mulched in Spring 2018 (treatment B) had the fewest weeds compared to beneficial habitat plants. You’ll also notice that in May there were still relatively few weeds in the plots where we tried to deplete the weed seed bank in the soil through solarization (treatment F) or repeated herbicide and tillage (treatment G).
What some of the plots looked like on May 16, 2019 when Bryan did the weed assessment.
By September 2019, the spring transplant treatments looked even better. Our wildflowers grew well during 2019 (with the help of some extra hand weeding). The plants we transplanted in Fall 2018 are still struggling and not nearly as large as the wildflowers in treatments A and B. I think this may have more to do with the weed competition they experienced that first fall (when we couldn’t plant for a few weeks after the buckwheat was mowed) than transplant timing. Hopefully they will catch up.
What some of the plots looked like on September 19, 2019 when Bryan did the weed assessment.
There are still a lot of weeds in the direct-seeded treatments (C, F, or G). Remember that our weed management strategy in these plots is repeated mowing to control annual weeds. Over time, the perennial wildflowers and grasses should take over. But it’s not supposed to be a quick method.
Spring transplant treatments (A and B) looked the best after their second full growing season. The fall transplants (D) had more weeds, but these plants also have been in the ground for one less growing season. I’m still hoping they will catch up. Each bar shows the average of four plots per treatment, and has an error bar showing variation among these plots (one standard error above or below the average for the treatment).
Effort
Most of the treatments we are comparing required much less work in their second year (2019) than in their first (2018). The exception is that we spent a lot more time hand weeding treatment D (buckwheat cover crop followed by fall transplanting) in 2019. Although we weeded the two spring transplanted plots the same number of times in 2019 (twice), it took longer to hand weed the plots without mulch. I’m not surprised. If you’re looking for the right establishment method for your project, you really need to ask yourself how much help you have available and when. If you can get a lot of people excited about helping you install the planting, but worry about getting consistent volunteers year after year, mulch may be the right choice for you. In the direct seeded treatments (C – spring; F – fall following solarization; G – fall following herbicide and tillage), the time input for 2019 was mowing, which was relatively quick. And we did just a little hand weeding of perennial weeds.
Transplanting (treatments A, B, and D) still takes more time than direct seeding, but the extra time we spent mulching in 2018 paid off in 2019 when we spent less time hand weeding (treatment B compared to A and D).
Costs
Nearly all of our costs were incurred in the first year of the project (2018). The only additional costs from 2019 were for gas to run the mower. We did replace a few plants in transplanted plots in Fall 2019, but we used some extra plants we had purchased in 2018. Below is the total cost of the plants and other supplies for each treatment. Transplanting will always be more expensive than direct seeding.
Treatment
Costs
A
$417.12
B
$539.29
C
$18.83
D
$390.55
E
$3.40
F
$149.10
G
$23.12
You may remember that we were also collecting insects. I promise I will write more about the insects we caught in another post. If you are tired of looking at snow and bare trees outside, you can see pictures of some of the insects we caught in my post from August 2019.
This post was written by Amara Dunn. All pictures were taken by her, unless otherwise credited.
This work is supported by:
Crop Protection and Pest Management -Extension Implementation Program Area grant no. 2017-70006-27142/project accession no. 1014000, from the USDA National Institute of Food and Agriculture.
New York State Department of Agriculture and Markets
Lots of flowers bloomed in 2019 in our habitat plots. Some we had planted, and some we hadn’t.
Obviously, nothing is growing right now, but I thought this would be a good time to update you on the success of our beneficial insect habitat plots during the 2019 growing season.
When things bloomed
Recall that the goal is to have at least one plant blooming all season long. We choose wildflower species accordingly, and it worked! The following table shows which months each species bloomed in 2019 (at least in the transplanted plots). An ‘X’ means the species was blooming during that month.
Wildflower
May
June
July
Aug
Sep
Golden alexanders
X
X
Catmint
X
X
X
X
Lanced-leaved coreopsis
X
X
Tall white beard tongue
X
X
Ohio spiderwort
X
X
Anise hyssop
X
X
X
Echinacea
X
X
X
Orange coneflower
X
X
X
Boneset
X
X
X
Wild bergamot
X
X
Common milkweed
X
NY ironweed
X
X
Showy goldenrod
X
New England aster
X
Blue false indigo
You may notice that the blue false indigo never bloomed in 2019, which was disappointing. Most of these plants are still alive (as you’ll see later in this post). They just didn’t bloom. Maybe next year?
Here’s what each species looks like:
Golden alexanders (Zizia aurea)Catmint (Nepeta faassinii)Coreopsis (Coreopsis lanceolata)Tall white beard tongue (Penstemon digitalis). Obviously not all of these flowers are white!Ohio spiderwort (Tradescantia ohiensis)Anise hyssop (Agastache foeniculum)Echinacea (Echinacea purpurea)Orange coneflower (Rudbeckia fulgida var. fulgida)Boneset (Eupatorium perfoliatum)Wild bergamot (Monarda fistulosa)Common milkweed (Asclepias syriaca)NY Ironweed (Vernonia noveboracensis)Showy goldenrod (Solidago speciosa)New England aster (Symphyotrichum novae-angliae)Blue false indigo (Baptisia australis); hasn’t bloomed yet in our field. Photo credit: Ansel Oommen, Bugwood.org
Transplanted wildflowers
The wildflowers in our transplanted plots are surviving pretty well (>80%). In the plots that were transplanted in the fall after the buckwheat cover crop, the survival is a bit lower. I think this has to do with some weed control issues (more on this in a future post).
How well have the transplanted wildflowers survived so far? Treatment A was transplanted in Spring 2018 and not mulched. Treatment B was transplanted in Spring 2018 and mulched. Treatment D was transplanted in Fall 2018 following a buckwheat cover crop. All have been hand weeded periodically. In both Spring and Fall of 2019 I counted plants to see how well they survived. The black lines on each bar in the graph show one standard error above and below the mean percent survival.
Some species have survived better than others, as the following chart shows. Again, we counted plants both in Spring and Fall 2019.
Do some species of wildflowers survive better when transplanted? There’s a little bit of variability, but overall most are surviving pretty well.
What about the direct-seeded plots?
Only three species of wildflowers planted by seed in Spring or Fall 2018 bloomed during the 2019 season. The table below shows which months these blooms were seen (marked with an ‘X’).
Common name
May
June
July
Aug
Sep
Coreopsis
X
X
X
Blackeyed susan
X
X
X
Partridge pea
X
X
Here’s what the flowers of blackeyed susan look like. The plant has much hairier leaves than the orange coneflower.
Blackeyed susan (Rudbeckia hirta)
And here’s the partridge pea:
Partridge pea (Chamaecrista fasciculata)
But, I also spotted some wild bergamot, tall white beard tongue, asters, golden alexanders, and either echinacea or orange coneflower seedlings. (I haven’t honed my horticultural skills enough yet to distinguish the foliage of these last two wildflowers.)
Seedlings of some wildflowers could be identified in the direct-seeded plots by September 2019.
There were also plenty of weeds blooming throughout the summer, and many of them were providing pollen and nectar for pollinators and natural enemies. Here are just a few examples:
From left to right: A bee feeding on a weed in the aster family, blooming chamomile, a lady beetle on a dandelion, and a bumble bee visiting clover (that wasn’t planted).
This table summarizes when during the season different weeds were in bloom. Again, an ‘X’ indicates the weed was blooming that month.
Weed
May
June
July
Aug
Sep
Campion
X
X
X
X
X
Chamomile
X
X
X
X
X
Clover
X
X
X
X
X
Dandelion
X
X
X
X
X
Vetch
X
X
X
X
X
Viola
X
X
X
X
X
Mustard
X
X
X
X
Deadnettle
X
X
Baby blue eyes
X
Henbit
X
Asters
X
X
X
X
Buckwheat
X
X
X
X
Oxalis
X
X
X
X
Plantain
X
X
X
X
Wild lettuce
X
X
X
X
Cinquefoil
X
X
X
Indian hemp
X
X
X
Redshank
X
X
X
Chickweed
X
X
Galinsoga
X
X
Geraniums
X
Sandwort
X
Grass
X
X
Horse weed
X
X
Lambsquarters
X
X
Ragweed
X
X
Black bindweed
X
Chicory
X
There’s more!
In addition to keeping track of what bloomed from May through September, we were also still tracking costs and time spent on each plot in 2019. And of course we collected a LOT of insects. But those stories will have to wait for another post.
This post was written by Amara Dunn. All pictures were taken by her, unless otherwise credited.
This work is supported by:
Crop Protection and Pest Management -Extension Implementation Program Area grant no. 2017-70006-27142/project accession no. 1014000, from the USDA National Institute of Food and Agriculture.
New York State Department of Agriculture and Markets
A tiny wasp lays its eggs in the larvae of the invasive lily leaf beetle. Releasing these wasps in the northeastern U.S. is a form of classical biocontrol. Photo credit: Dan Gilrein
Last week, some colleagues told me about a cool online database that will help you learn more about how classical biocontrol is helping us fight invasive insects.
Classical biocontrol means introducing a natural enemy of a pest to help manage that pest. The natural enemy establishes a population where you have released it (and maybe even spreads), so that you don’t need to repeatedly release more natural enemies. It is a strategy that can be especially useful against invasive pests.
One thing that makes a pest invasive is the fact that when it arrives in a new place (for example, on a new continent), native organisms don’t eat it because they have not evolved with this new pest as a food source. Sometimes scientists can search the geographic area from which the invasive pest came and find a natural enemy of that pest. Many tests are done over a long period of time in order to assess potential unintended consequences of introducing this natural enemy to a new place. For example, scientists determine whether the new natural enemy is likely to also impact populations of native organisms (especially those that are not pests). Only after extensive study will this new natural enemy be released to help reduce populations of the invasive pest.
When done carefully, classical biocontrol can be a lower-risk solution to managing invasive pests compared to chemical pest management. It is also a long-term solution. The new natural enemy reproduces in its new geographic range and brings the invasive pest population into balance. The invasive species won’t be eliminated, but it will likely do less damage.
A new database from the University of Massachusetts lets you learn more about insects that have been introduced to North America to control invasive insect pests as classical biological control agents. You can Search the Catalog by the scientific name of the target pest, the scientific name of the natural enemy, information about where and when the natural enemy was first released, or other criteria.
The invasive hemlock woolly adelgid on a hemlock branch. Several different classical biocontrol agents have been released in the U.S. to manage this invasive pest. Photo credit: Amara Dunn
You will need to know the scientific name of the pest or natural enemy you are interested in, but a quick Google search can help you with that. For example, Adelges tsugae is the hemlock woolly adelgid, which you may have heard about. If not, you can learn more here. Laricobius nigrinus was released to help manage hemlock woolly adelgid. Other examples include Agrilus planipennis (emerald ash borer) and Lilioceris lilii (lily leaf beetle). NYS IPM is involved in a project to use classical biocontrol to manage this last pest in NY.
As the days start to get shorter and cooler, you might find yourself spending more time indoors. And if that’s the case, why not spend some time learning more about how classical biocontrol is helping to manage pests in the landscapes around you?
You’ve read about all the different methods we are testing for establishing native wildflowers and grasses as habitat for pollinators and natural enemies of pests. You know we learned a lot in our first season. You know we’ve been using several different techniques to collect insects in these plots. And you saw a pictorial summary of our sampling and some of the insects we’ve caught in Summer 2019.
Wouldn’t you like to come see these plots in person, hear about our preliminary results, and learn more about attracting pollinators and other beneficial insects to your farm or yard?
If you live reasonably close to Geneva, NY, you can! We are having two field events this fall:
On Wednesday, September 25, 2019, stop by our field between 3:30 and 6:30 PM for an Open House. There will be no program, just stop by and talk with Betsy Lamb, Brian Eshenaur, and I. All the details can be found here, including the address and a map to help you find our field.
On Thursday, September 26, 2019, we have a Twilight Field Day from 5 to 7 PM. This meeting has been planned with growers in mind (especially Christmas tree and nursery growers). DEC credits (1.5) will be available for categories 1a, 3a, 24, 25, and 10, and dinner is included. The cost for this meeting is $15, and we need you to register so we know how much food to provide. All the details (including the registration link) can be found here.
If you’re coming to either of these events, we’ll have lots of signs up to help you find our field. Look for the following image:
This work is supported by:
Crop Protection and Pest Management -Extension Implementation Program Area grant no. 2017-70006-27142/project accession no. 1014000, from the USDA National Institute of Food and Agriculture.
New York State Department of Agriculture and Markets
Summer isn’t over yet for farmers and extension staff doing field experiments!
Labor Day weekend may be viewed by some as the end of summer, but farmers know that the summer growing (and harvesting!) season is far from over. Similarly, the field projects I’m involved with this summer (read more here and here) are still running. Over the fall and winter I’ll be analyzing data and sharing results (on this blog, and at winter meetings). In the meantime, here’s a pictorial summary of my summer projects (so far).
Golden alexanders (Zizia aurea) was our earliest-blooming wildflower in our beneficial insect habitat plots around the Christmas trees. It was blooming on May 16 in Geneva, NY.Some of us may not like them in our lawns, but starting in the first week of May (Geneva, NY) dandelions were providing food for beneficial insects like this bee.This shy lady beetle was finding food in the flowers of this weedy mustard plant in mid-May (Geneva, NY).After expert training from Cornell entomologist Jason Dombroskie (pictured here during our training session in late April), we’ve been using a sweep net to catch insects that fly or perch on the wildflowers in the habitat plots we started last summer. We sampled this way once every month.Starting the week of May 20th, we set out pan traps (blue and yellow plastic bowls filled with soapy water and weighed down with rocks) approximately every other week. These traps catch insects flying through our plots, especially those that are attracted to the colors blue and yellow. This includes many bee species.Also during the week of May 20th, we started setting pitfall traps once each month. Insects walking along the ground fall into these deli cups filled with a drowning solution. We put rain covers over them (made out of clear plastic dinner plates and wire from old flags) to prevent a heavy rain from flooding the deli cups during the 3 days the traps are set.We caught and saw so many insects (and non-insects, like spiders) this summer! This seven-spotted lady beetle was a frequent visitor to our plots.Many flies are important pollinators, like this one that resembles a bee at first glance. Many flies are also important natural enemies of pests (either as adults, or as worm-like larvae).This minute pirate bug may be tiny (it’s magnified 20X), but it is an important natural enemy of pests.We caught so many different kinds of bees and wasps!Plenty of caterpillars (like these monarchs) enjoyed munching on the foliage of our wildflowers.And in late July, we started seeing adult butterflies visiting the flowers like the viceroy butterfly on these purple coneflowers (Echinacea purpurea).I learned that these are tiger beetles. They are fast-moving ground predators, and we caught a lot in our pitfall traps.Different wildflowers bloomed at different times, like these purple catmint (Nepeta faassinii) and tall white beard tongue (Penstemon digitalis) in June.White boneset (Eupatorium perfoliatum) and pale purple wild bergamot (Monarda fistulosa) were blooming in late July.And now the rudbeckia (two different species, but Rudbeckia fulgida var. fulgida is pictured here) and deep magenta NY ironweed (Vernonia noveboracensis) are in full bloom. But the asters and goldenrod haven’t started, yet.And the Christmas trees planted around these beneficial insect habitat plots keep growing!I couldn’t have done this without the help of my great co-workers, Betsy Lamb, Deb Marvin, and Brian Eshenaur! They were still smiling after a morning of weeding the wildflowers by hand!A student from a local college helped me a lot with insect collection!Meanwhile, field trials with biofungicides are ongoing, targeting cucurbit powdery mildew on winter squash and white mold on snap beans and tomatoes (not pictured). This project is funded by the New York Farm Viability Institute.Elizabeth Buck (left) and Crystal Stewart (right) are running the trials in western NY and eastern NY, respectively. This project is funded by the New York Farm Viability Institute.Meg McGrath (left) is running the trial on Long Island, but we all got together at a twilight meeting in eastern NY last week. This project is funded by the New York Farm Viability Institute.So far, it’s been a good summer! I’ve really enjoyed working with great colleagues and learning new things!
The field projects I’ve just described will be wrapping up in September. Check back to learn about the results. Better yet, click the green “Subscribe” button towards the top and right of this page, and you’ll receive an email when a new post is available!
In the meantime, there will still be at least a few more weeks of pictures posted regularly on Twitter (@AmaraDunn) and Instagram (@biocontrol.nysipm).
Creating habitat for beneficial arthropods was supported by:
Crop Protection and Pest Management -Extension Implementation Program Area grant no. 2017-70006-27142/project accession no. 1014000, from the USDA National Institute of Food and Agriculture.
New York State Department of Agriculture and Markets
This month’s post is about a project being led by Jaime Cummings, the Field Crops and Livestock IPM Coordinator at NYS IPM. The goal is to improve biological control of the cereal leaf beetle, a pest of small grains. Before we tell you about the biocontrol project, you’ll need some background information on this pest and the other management options available. You can use the following links to navigate to each section of this post:
Cereal leaf beetle damage on the flag leaf of a winter wheat plant. (Photo by J. Cummings, NYS IPM)
Cereal leaf beetles and the damage they cause
The cereal leaf beetle (CLB), Oulema melanopus, can be a significant pest of winter and spring small grains production in NY, especially in parts of western NY. This invasive species was first detected in Michigan in 1962, and has since become established in many grain producing states in the US, despite quarantine and pesticide eradication efforts in the 1960’s and 1970’s.
Figure 1. Cereal leaf beetle adult (A) and larval (B) stages. (Photos by J. Cummings, NYS IPM)
You may be familiar with this pest either in the larval or beetle stage (Fig. 1). CLB has one or two generations per growing season, and the adults overwinter in hedgerows, woods or field margins. We usually start seeing the adults move into small grains fields in April or May to lay eggs which develop into the damaging larvae. The larger the larvae get, the more damage they inflict on the crop. After about two weeks of feeding, the larvae drop to the ground and pupate for about two weeks before the adults emerge again.
When looking for these pests, keep an eye out for the typical larval feeding damage that looks like strips of green tissue missing between leaf veins. Severely damaged leaves may appear skeletonized, and intense feeding pressure in a field may result in a ‘frosted’ appearance of flag leaves (Fig. 2).
Figure 2. Severe cereal leaf beetle larval feeding on winter wheat. (Photo by J. Cummings, NYS IPM)
Considering that the top two leaves of the wheat/barley/oat crop are what contributes most to grain yield, severe infestations of CLB can significantly impact yield and grain quality. Even in small grain or mixed stand forage crops, this pest can have negative effects on the yield and quality of the forage because they can significantly reduce leaf area and photosynthetic capability of the crop.
Scouting for cereal leaf beetle and deciding when to spray
It’s important to scout for this pest, usually starting in early to mid-June when larvae are first appearing. The economic threshold for insecticide application for CLB is when you count an average of three or more larvae per plant before the boot stage or one or more larvae per flag leaf after the boot stage. Occurrence of this pest can be inconsistent within a field, therefore plan to scout weekly and walk a random pattern throughout each field stopping at 10 random locations to count larvae on 10 plants at each location. Because insecticides labeled for CLB target the larval stages, in order for your pesticide applications to be most effective, make sure that at least 25% of CLB eggs have hatched and that larvae are present and actively feeding when you decided to spray. And, if you’re seeing adults in late June or beyond, it’s probably too late to spray for the larvae. (Always follow label recommendations and restrictions when applying pesticides)
Paying attention to CLB populations in your fields via scouting is an important part of an integrated management approach for minimizing losses to this pest. A growing degree day (GDD) model for CLB developed in Michigan determined that adult CLB begin to emerge around 350-400 GDD (base 48) to begin egg laying.
Biocontrol of cereal leaf beetle
Unfortunately, there is no specific host plant resistance available for CLB, but there are natural predators of the larvae and eggs which can help to keep the pest population in check, and possibly below the economic threshold when well-established in an area. Lady beetles are known to prey on CLB larvae and eggs, and there is at least one egg parasite though it is not widely distributed.
There is also a CLB larval parasitoid wasp, Tetrastichus julis, which was originally introduced from Europe as a biological control agent in Michigan in 1967 (Fig. 3). Subsequent releases into other states, including NY in 1973, have led to a sporadic establishment of this biological control parasitoid throughout small grain production areas of the US.
Figure 3. Tetrastichus julis, a parasitic wasp on a cereal leaf beetle larva. (Photo courtesy of Washington State Department of Agriculture)
Our project: Improving biocontrol of cereal leaf beetle
Given that CLB damage can be widespread and undermanaged in many small grains fields in NYS, and under the advice of Dr. Elson Shields (Cornell University Field Crops Entomologist), the NYS IPM program decided to try to determine the parasitism levels of CLB larvae in various locations around the state and to try to increase populations of the parasitoid in the Aurora area of Cayuga County, where the CLB tends to be a perennial pest. The multiyear project was initiated this year, with CLB larval collections from locations in six counties. However, there were no CLB present to collect at two of the locations, so the data collected in 2019 includes only four locations (Table 1).
Table 1. Cereal leaf beetle collection efforts for determining parasitism levels in 2019.
Location
County
Collection date
Crop
# CLB larvae collected
Seneca Falls
Seneca
6-Jun
winter wheat, rye, barley
96
Aurora/Musgrave
Cayuga
12-Jun
spring barley
92
Ithaca
Tompkins
12-Jun
winter wheat, rye, barley
45
Penn Yan
Yates
13-Jun
spring oats and peas
110
Oriskany
Oneida
11-Jun
winter wheat
0
Homer
Cortland
10-Jun
winter wheat
0
At each location, a target of approximately 100 CLB larvae of all different sizes/growth stages were collected by hand from wheat, barley or oat fields. The larvae were temporarily reared in incubation chambers on host plant leaves until approximately half of the larvae were dissected to determine baseline parasitism levels for each location (Fig. 4).
Figure 4. Cereal leaf beetle rearing chambers (A) and dissection process (B). (Photo by J. Cummings, NYS IPM)
The eggs of the parasitoid are visible when the CLB larvae are cut open under a microscope (Fig. 5).
Figure 5. Dissected CLB larvae, and one with T. julis parasitoid eggs. (Photo by J. Cummings, NYS IPM)
After baseline parasitism levels were determined for each collection location, the other half of the CLB larvae were then released at the Cornell Musgrave research farm near Aurora, NY (Fig. 6). This process will be repeated over the next few years.
Figure 6. Cereal leaf beetle larvae with known level of parasitism being released in Cayuga County (Photos by J. Thomas-Murphy, Cornell University)
The goals of this project are to determine the established levels of the T. julis parasitoid around the state since the initial release in 1973, and to try to determine if we can increase its population at the research farm through consecutive releases. From this first year of data collection, we know that the parasitoid population is low at the research farm in Cayuga County (6%) and at two of the collection sites (7% and 10%, in Seneca and Yates Counties, respectively), but was at approximately 30% at the Ithaca (Tompkins County) collection site (Fig. 7).
Figure 7. Percent T. julis parasitized cereal leaf beetle larvae collected from various locations.
We also know that although there has been a need to spray insecticides to manage CLB at the research farm in Cayuga County and near the other collection sites, there has been no need to spray for CLB at the Ithaca (Tompkins County) collection sites. It’s likely that the T. julis parasitoid population at the Ithaca site keeps the CLB population below economic threshold levels. We hope that by intentionally distributing this parasitoid into an area with known CLB problems, we can establish a robust parasitoid population that may result in a reduction of necessary insecticide sprays for this pest.
This post was written by Jaime Cummings, Ken Wise, and Amara Dunn, all of the New York State Integrated Pest Management Program.
Caption: Lady beetles will eat any aphid species, but other aphid natural enemies are much more selective. (Photo credit: Amara Dunn)
Practicing good integrated pest management in the greenhouse requires correct identification of the pest. Accurate pest ID is also critical to successful use of biocontrol. Aphids are a good example. Biocontrol of aphids works best when you match the biocontrol agent to the aphid species you have. When I first learned this, I was a bit intimidated, because aphids are pretty small, and I’m not an entomologist. But the four aphid species you are most likely to encounter in your greenhouse are actually pretty easy to differentiate.
Anatomy of an aphid
In order to successfully ID aphids, you need to know (just a little) about aphid anatomy. All aphids are pretty small (between approximately 1/16 and 1/8 inches long). In addition to six legs and a body, aphids have antennae. Antennae attach near their eyes and are angled back over their bodies. They also have two little “spikes” that protrude from their rear end. These are called cornicles. Not so bad, right?
Two features that will help you identify an aphid are the antennae attached to their head, and the two short cornicles attached to the rear end of their abdomen. (Diagram credit: Amara Dunn)
Green peach aphid
Green peach aphids vary in color from green to pink. Between their antennae you’ll see an indentation, and their cornicles are the same color as their body, with dark tips. (Photo credit: John Sanderson)
Green peach aphids come in different colors (from green to, well, peachy pink) and they are one of the smaller species. Their cornicles are the same color as their body (whatever that color is), and have dark tips on the ends. Green peach aphids also have an indentation in their head between the bases of their antennae.
Melon (or cotton) aphid
Melon (also called cotton) aphids can be distinguished from green peach aphids by their dark cornicles. They also lack an indentation between their antennae. (Photo credit: John Sanderson)
Melon aphids (also called cotton aphids) also come in a range of colors that include light yellow, green, dark green, or almost black. Regardless of the body color, the cornicles will always be dark. Also, there’s no indentation in their head between the bases of the antennae. This is another small aphid species.
Foxglove aphid
Foxglove aphids are shiny green with long antennae that look like they have dark spots on them. You’ll also see darker green spots at the base of each cornicle and dark leg joints. (Photo credit: Dan Gilrein)
Foxglove aphids are large (for an aphid). Their bodies are light green, but often shiny. There is an indentation in their head between their antennae. Their antennae are extra-long, extending well beyond the end of their body, and appear to have dark spots on them because the joints of the antennae are dark. The joints of their legs are also dark. Check where the cornicles attach to the body of the aphid. Foxglove aphids have darker green spots on their bodies at the base of the cornicles. These aphids usually like to hang out on the lower leaves of a plant, though they will infest flower petals sometimes.
Potato aphid
Potato aphids have a dark stripe running down the length of their body, and they look faintly segmented. (Photo credit: John Sanderson)
Another large aphid, potato aphids come in pink and green. They look like they have a dark stripe running down the middle of their backs, and their body appears faintly segmented. They also have an indentation in their head between the antennae. Of the four species we’re discussing here, only the melon aphids lack this indentation.
To see these features, you will need a little magnification, but you don’t need a fancy microscope. Find a hand lens or a magnifier with 10X magnification. I like to keep one in my backpack so I’m always prepared.
A 10X hand lens will enable you to magnify the features of an aphid that are important for identification.
There are even some relatively inexpensive 10X lenses you can snap on to your smartphone or tablet. Not only does this turn your device into a little microscope, but you can take a picture to document what you see (and show to an expert, later).
Magnifying macro lenses can be clipped onto your smart phone or tablet, helping you both magnify and document the aphids you find.
You can also find (at least some of) these four aphid species outside. Last summer I spotted the aphid below on an acorn squash plant in August. Now that you know what to look for, what species do you think it might be?
Use what you’ve learned to identify this aphid! (Photo credit: Amara Dunn)
One minor complication: Each of these four aphid species can either have wings, or be without wings. Usually aphids you find in a greenhouse have no wings, so you can stick with the above descriptions. But winged aphids can appear in the greenhouse, particularly when populations get very high. If you find aphids with wings in your greenhouse, the above descriptions won’t apply; ask for some help from your local extension office.
Both of the aphids in this picture are green peach aphids, but the one on the right has wings, and would be tricky to identify using the criteria described in this post. Get some help from an expert. (Photo credit: Whitney Cranshaw, Colorado State University, Bugwood.org)
Choosing the right natural enemy
An Aphidius parasitoid wasp lays an egg inside its aphid prey. The developing wasp will kill the aphid. These happen to be cabbage aphids. (Photo credit: David Cappaert, Bugwood.org)
A good biocontrol option for aphids is a parasitoid wasp from the genus Aphidius. These tiny wasps are called parasitoids because they lay their eggs inside of aphids. As the young wasp grows, it kills the aphid and turns it into a mummy.
Aphids that are eaten from the inside out turn into dry, brown “mummies”. On this aphid mummy you can see the hole from which the adult wasp emerged. (Photo credit: Ken Wise, NYS IPM)
But if you want to purchase Aphidius wasps to release in your greenhouse (or the banker plants and prey that support them; read more here), you’ll need to know which Aphidius species to use. Aphidius colemani works well against green peach and melon aphids, while Aphidius ervi works well against foxglove and potato aphids. Another natural enemy you can use is Aphidoletes aphidimyza. This is a tiny fly whose larvae are voracious aphid predators. Although it seems to be less effective against foxglove aphid, it may work well in combination with another natural enemy.
The larvae of the tiny fly Aphidoletes aphidomyza crawls around on leaves searching for aphids to eat. (Photo credit: Sarah Jandricic)
Like all biocontrols, Aphidius wasps and Aphidoletes larvae need to be released while your aphid population is very small, before it gets out of hand. Aphid infestations can explode very quickly! Scout your crop regularly, and keep records so you know which aphid species you are likely to have. (Consider the Pocket IPM Greenhouse Scout app to help you with your scouting and pest management.) Then plan your biocontrol releases accordingly. Parasitoids and predators for aphids should be released preventatively on crops that are prone to aphids.
If you’ve inspected your aphids at 10X magnification, and still aren’t sure which species you have, contact your local extension office for help with ID. If you are planning to send a picture, make sure that it is clear and shows the features of the aphid that you now know are important (antennae, body, cornicles).
You can learn more about aphid biocontrol in this factsheet from John Sanderson (Department of Entomology, Cornell University) on managing aphids in a greenhouse. Identification of these four common aphid species and which biocontrols you can use against them are also summarized here. The natural enemies listed in the chart are meant to be a starting place. Maximizing the efficacy of your aphid biocontrol program takes some trial and error and willingness to fine-tune your program to the crop and environmental conditions you’re dealing with. Suppliers of aphid natural enemies also have great information about how to use these biocontrol agents most effectively.
Mulching transplants planted in Spring 2018 was the most expensive establishment method, but these plots were looking pretty good a year later, even before we weeded.
By the end of our first field season, we had started using six different methods to establish wildflowers as habitat for beneficial insects (plus a weedy mowed control treatment). We also collected data on how much time and money we spent on establishment and how successful our weed management was. You can read about results from Year 1 in my post from last November.
But beneficial insect habitat establishment is not a one-year project. The establishment methods we started to implement in 2018 are ongoing, including periodic mowing of direct seeded plots, and hand-weeding of transplanted plots. We’ll keep track of how much time and money we invest in these plots in 2019, too.
Plots that were direct seeded in 2018 will be mowed this year to favor the perennial beneficial insect habitat plants over annual weeds. This plot was treated with alternating herbicide and tillage during Summer 2018, and wildflower seed was planted in Fall 2018; (A, left) plot before mowing, (B, right) same plot after mowing.Plots that were transplanted in 2018 will be hand weeded this year to help the perennial wildflowers and grasses out-compete weeds. This plot was transplanted in Spring 2018 into bare (not tilled) ground and no mulch was used; (A, left) plot before weeding, (B, right) same plot after weeding.
And we want to know whether these plots are actually attracting beneficial or pest insects. So, in 2019 we are starting “Phase II” of our beneficial insect habitat work. We want to know which and how many insects (and other arthropods, like spiders) are being attracted to each type of plot. We will also count insects in no habitat plots (weedy, mowed occasionally) and mowed grass plots in the middle of the Christmas tree field for comparison.
Insect collection began in early May, and we are using four different techniques:
Sweep net – This is what it sounds like. We “sweep” a net through the air above the ground to capture mostly flying insects, or those who may be resting on the plants.
Butterfly and moth count – We walk through the field, counting how many of each butterfly or moth species we see in each plot.
Pan traps – These are bright yellow and blue bowls filled with soapy water. One bowl of each color is placed in each plot for 2 days, then we collect the insects that have been attracted to the colorful bowls and were trapped in the soapy water. This method will help us count flying insects, especially bees and wasps.
Bright blue and yellow bowls filled with soapy water and weighed down with rocks will attract certain flying insects. By counting insects collected in these pan traps, we can learn which insects are spending time in each plot.
Pitfall traps – These are clear plastic 16-oz deli cups (like you might use for take-out food) that are sunk into the ground in each plot. Insects that crawl along the ground fall in. We will use this method to count mostly ground-dwelling insects.
A pitfall trap collects ground-dwelling insects. This one is protected by a rain cover. We didn’t want all the rain we’ve been getting this spring to overflow the traps and wash away the insects we caught!
I will write another blog post or two about this project during or at the end of this season. If you want to see more frequent updates, follow me on Twitter (@AmaraDunn). I’ll post weekly pictures of this project, including which beneficial insect habitat plants are blooming each week. You can also see lots of pictures from this project on Instagram (biocontrol.nysipm).
This work is supported by:
Crop Protection and Pest Management -Extension Implementation Program Area grant no. 2017-70006-27142/project accession no. 1014000, from the USDA National Institute of Food and Agriculture.
New York State Department of Agriculture and Markets
Many crops (and plenty of non-crop plants) rely on pollinators. Let’s protect them!
As I’ve discussed before, the natural enemies that provide biological control of pests include both larger creatures (like insects, mites, and nematodes) and microorganisms (fungi, bacteria, and viruses) that combat pests in a variety of ways. Microorganism natural enemies are regulated as pesticides (one type of biopesticide), while the larger natural enemies are not. Growers who are successfully using biocontrol insects, mites, and nematodes usually recognize that they need to apply pesticides in such a way that they are compatible with the biocontrol organisms they use. Take a look at my April post for a summary of online resources that can help you check compatibility of pesticides (including biopesticides) with natural enemies.
Some of these compatibility resources include information on the effects of pesticides (and biopesticides) on bees. Pollinators (including honey bees, lots of other bees, and some non-bees) are very important beneficial insects. You may have noticed that they have found their way into several of my blog posts. So, I wanted to let you know about a brand new resource (hot off the digital presses) to help you protect pollinators.
“A Pesticide Decision-Making Guide to Protect Pollinators in Tree Fruit Orchards” is a terrific resource to help you choose pesticides (and pesticide combinations) that are least-toxic to bees.
“A Pesticide Decision-Making Guide to Protect Pollinators in Tree Fruit Orchards” was written by Maria van Dyke, Emma Mullen, Dan Wixted, and Scott McArt. Although it’s focus is tree fruit orchards (and therefore the pesticides used in them), it should be useful for growers of other crops who want to choose pesticides that are least toxic to bees. A few highlights:
It includes information not only on pesticides used alone, but (when available) on synergistic effects when multiple pesticide active ingredients are used together. When you combine some chemicals (either in the tank or in the environment) the mixture is more toxic than both chemicals alone.
Where available, it summarizes pesticide toxicity to other bees besides just honey bees (e.g., bumble bees and solitary bees). You can read more about why this is important in this recent article.
It describes what we know about sub-lethal (in other words, negative effects on the bees that are less serious than death) effects of pesticides on bees.
It includes about half a dozen biopesticide active ingredients.
Pollination is being done by more than just honey bees! This bumble bee (plus many more bee species) are important pollinators in NY.
Guides for other crops and other resources for growers wanting to protect pollinators can be found here.
You might be asking: If a chemical on this table is toxic to bees, will it also be toxic to the insect and mite natural enemies I am releasing or conserving on my farm or in my garden? I wish I had a definitive answer to that. As you can see from the nearly three pages of Literature Cited at the end of this document, collecting these data is a time-consuming process. For now, stick with the compatibility resources that are already available, and ask the companies you buy from (pesticides or natural enemies) about compatibility.
In closing, a huge amount of work went into this resource to summarize so much useful and current (as of October 2018) information in an easy-to-read table. Bravo to the authors! The Pollinator Network @ Cornell has other helpful resources for growers on protecting pollinators. Winter is a great time to make plans for using IPM and protecting the pollinators and natural enemies that are so good for the crops we grow!
Fair warning, this is going to be a longer post. But partly that’s because there are so many pictures. I will start with the overview, then go a bit deeper into the weeds (literally and figuratively). To help you navigate more quickly, here’s a sort of table of contents that will quickly take you to the information you may be most interested to read:
Some of our beneficial insect habitat plots looked really beautiful this fall! Others are still works in progress.
Overview
Remember back in June when I told you about the different techniques we were comparing for establishing habitat for beneficial insects? Time for an update! Here’s a brief, two-page summary of the first year of this project. For all the juicy details (and lots of pictures), keep reading!
First, remember that when I say “beneficial insects”, I mean both pollinators and natural enemies of pests. (Technically, arthropod would be a better term than insect, because spiders and predatory mites are some of the beneficial creatures we’d like to attract.) Fortunately, the same type of plants provide food and shelter for both pollinators and natural enemies on your farm or in your garden.
We used six different techniques to establish this habitat during Spring, Summer, and Fall of 2018. Treatment E was our control, where we did nothing but mow (after initial herbicide applications).
Treatment
Fall 2017
Spring 2018
Summer 2018
Fall 2018
A
Herbicide
Herbicide, transplant
Weed 2x
Replace dead plants
B
Herbicide
Till, transplant, mulch
Weed 2x
Replace dead plants
C
Herbicide
Till, direct seed
Mow 3x
Mow 1x
D
Herbicide
Till, plant buckwheat
Mow 1x, till, plant buckwheat
Mow 1x, transplant
E – control
Herbicide
Herbicide
Mow 3x
Mow 1x
F
Herbicide
Till, lay plastic
Continue solarization
Remove plastic, direct seed
G
Herbicide
Herbicide/till
Herbicide 2x, till 1x
Till 1x, direct seed
We transplanted the following species in treatments A, B, and D:
Common name
Scientific name
Number of plants in each 5 x 23 ft plot
Anise hyssop
Agastache foeniculum
2
Common milkweed
Asclepias syriaca
3
Blue false indigo
Baptisia australis
2
Lanced-leaved coreopsis
Coreopsis lanceolata
3
Purple coneflower
Echinacea purpurea
2
Boneset
Eupatorium perfoliatum
3
Wild bergamot
Monarda fistulosa
2
Catmint
Nepeta faassinii
2
Tall white beard tongue
Penstemon digitalis
3
Black-eyed Susan
Rudbeckia fulgida va. Fulgida
1
Little bluestem (grass)
Schizachyrium scoparium
11
Showy goldenrod
Solidago speciosa
1
New England aster
Symphyotrichum novae- angliae
3
Ohio spiderwort
Tradescantia ohiensis
2
NY ironweed
Vernonia noveboracensis
2
Golden alexanders
Zizia aurea
3
We planted seeds in treatments C, F, and G. The seed mixture we used was the Showy Northeast Native Wildflower & Grass Mix from Ernst Seeds, which included a more diverse species mix. This mix changes a bit from year to year. If you’re interested, you can learn about the details of the specific mix we used here.
Labor and costs
Not surprisingly, there were big differences in how much time and money we spent on different treatments this first year. The costs and hours below are for a total area of 460 ft2 (0.01 A) per treatment. Most of the cost differences are due to the huge difference in seed versus transplant expenses. We paid about $2 per plant and needed 180 plants for each treatment. In contrast, we spent about $12.50 on seed for each treatment. You can find itemized lists of cost and time inputs for each treatment here.
Treatment
Supply costs
Time (person hrs)
A – spring transplant
$417.12
13.2
B – spring transplant & mulch
$539.29
20.4
C – spring seed
$17.75
4.4
D – buckwheat & fall seed
$390.55
10.3
E – control
$2.32
2.6
F – solarize & fall seed
$148.02
10.2
G – herbicide/tillage & fall seed
$22.04
6.3
But, there were also big differences in how quickly the plants established. By September, both treatments (A and B) that had been transplanted in the spring looked like well-established gardens, with large, blooming wildflowers.
Four and a half months after transplanting, the beneficial habitat plants in treatments A (left) and B (right) were mostly growing well. But there was a big difference in weed control, in spite of similar amounts of time spent weeding each treatment.
We were generally pleased by how well most of the spring transplants survived. Although all the transplants came in 50-cell flats, some were larger than others, and the larger transplants survived better. We were fortunate to be able to plant into nice moist ground, so except for a little water on the day of transplanting, we didn’t irrigate. Survival might not have been as good if we’d had different planting conditions.
In contrast, the much less expensive treatment C was not looking too impressive even by October. A few partridge peas and blackeyed Susans bloomed this year, but otherwise it didn’t look much different from the control plots. In mid-summer, it looked like we were growing more ragweed than wildflowers.
In late July, it looked like we were growing mostly ragweed in treatment C (left). But after mowing four times during the summer and fall, you could definitely see the blackeyed Susans establishing (right).
Two of the treatments (F and G) were planted with seeds this fall, and one treatment (D) was transplanted this fall. So it’s really too early to tell how successful those treatments were. Stay tuned for more updates!
Fall transplanted (D) or direct seeded treatments (F and G) did not look very impressive by October 19, 2018. I’m curious to see what they look like next spring, summer, and fall!
Details on weed control
What about weeds? The graph below shows the average percent of the surface area of each plot that was covered with weeds versus planted beneficial habitat species on September 19, 2018. (Thank you, Bryan Brown, NYS IPM Integrated Weed Management Specialist for doing a weed assessment for us!) While we spent about the same amount of time weeding treatments A and B (the time difference is due to the time spent mulching treatment B), we achieved much better weed control with the mulch than without it!
Bryan Brown assessed the percent of each plot that was covered with either the beneficial habitat species we had planted (blue) or weeds (orange). Each bar represents the average of four plots for each treatment, and the error bars show the standard error.
In treatment B, we spread chipped shrub willow mulch about 3 inches deep around the transplants. If I were to do this again, I would spread it thicker. I was disappointed with how many weeds were growing through the mulch just a month after transplanting.
The chipped shrub willow mulch we used was not as effective at suppressing weeds as I had hoped. On the left is part of the plot that had not been weeded yet. On the right is the part that was weeded on July 6. You can also see from this picture that there was a lot of lambsquarters in this field, and that we hadn’t been able to seed grass between the plots, yet.
But weeding twice during the season pretty much took care of the weeds in treatment B. Treatment A was also weeded twice, but as you saw in the graph earlier, weed control by the end of the season was not as effective.
Treatment A (transplanted in the spring, with no additional weed control) before (right) and after (left) hand weeding on July 6.
I think we’ll have to wait until next year to really understand how weed control is working in treatment C. Remember, the strategy was to slowly deplete the annual weed seedbank by allowing weeds to germinate, but preventing them from producing more seed. This is not supposed to be a quick establishment method, and it wasn’t.
This is what partridge pea looks like. It was one of the few species from the native wildflower and grass mix seeded in the spring (treatment C) that bloomed during this first growing season.
Buckwheat as a weed-smothering cover crop
By the time Bryan did our weed assessment, it had been 3 weeks since we mowed the second planting of buckwheat. Ideally, we would have transplanted shortly after mowing the buckwheat. But, the second crop of buckwheat was starting to set seed by the end of August, and our transplants weren’t scheduled to arrive until the end of September. So we mowed the buckwheat early to prevent it from contributing its own seed to the weed seedbank. But this meant that a lot of weeds had time to germinate before we transplanted the habitat plants. The buckwheat certainly suppressed a lot of weeds during the growing season, and I hope that this will help reduce weeds next year.
The buckwheat established quickly and crowded out many weeds. We mowed the first crop in July and re-planted. We had to mow the second crop about 3 weeks before we transplanted habitat plants (not ideal).
Solarization
Overall, we were pleased with how the solarization worked. We laid down 6 mil clear plastic (leftover from a nearby high tunnel) in early June, and did a little weed control around the edges of the plastic just once during the summer to prevent more weed seed production and to prevent shading of the plots.
We laid 6 mil clear plastic over some plots on June 5, 2018 to solarize the soil underneath (and kill weeds).
We also learned that solarization will not control purselane. In contrast, the purselane thrived only under our clear plastic, and nowhere else in the field. The plot that had the most purselane also had the most other (mostly grass) weeds. I think the purselane pushed the plastic away from the soil and reduced the temperature a bit, allowing other weeds to grow.
In some solarized plots, purselane grew happily under the plastic. Purselane was not a common weed anywhere else in the field during the season.
Some other plots were virtually weed-free when we pulled the plastic up in October. (Did you see how large the error bar was for weeds in treatment F in the weed graph above? This means there was a lot of variability between plots in this treatment.) Our soil temperature probe happened to be in the plot with the most purselane, and we still achieved maximum soil temperatures of 120 °F (at a depth of about 3 inches), compared to 90 °F in a nearby control (treatment E) plot.
Solarization results were pretty variable from plot to plot. In some plots, it worked great (left). In other plots more purselane germinated and it didn’t work as well (right). We cut all the weedy vegetation off at the ground before direct seeding the beneficial habitat plants.
Repeated herbicide and tillage
At the weed assessment in September, the plot that had been alternately treated with herbicide and tilled looked best in terms of weed control. Like treatment C and all the treatments planted (by seed or by transplant) in the fall, I think we’ll get a better idea next year of how effective this method was at suppressing weeds.
A few weeds were present a week after the last time the herbicide/tillage treatment (G) was rototilled. We broadcast, raked, and pressed beneficial habitat seed into these plots.
Timing of fall planting
One thing we struggled with this fall was deciding when to plant the wildflower and grass seed mixture. One source recommended the seeds be planted sometime between October and December. We were cautioned that if we planted the seed too early, some species (especially blackeyed Susans) might germinate this fall, and the young seedlings would be killed by an early frost before they established. But we were also afraid of waiting too long and not being able to till the soil (treatment G, only) if it got too wet. And we wanted a nice smooth seedbed. In treatment F, we suspected that leaving the clear plastic on into November would protect the weeds from the cooler weather. But we worried that taking it off too early would only allow more weed seeds to blow onto the bare ground.
We direct seeded October 18, 2018, after the weather cooled down a bit, and before the ground got too wet.
Finally, we compromised and planted the seeds on October 18 and 19, after our first hard frost, and once it looked like the nighttime temperatures would be in the 40’s (or below) for the next 10 days. It was only a week after the last tillage in treatment G, and the soil was still relatively dry. Those who live in the Finger Lakes know that late October and early November were pretty wet this year, so I’m glad we planted when we did. If you are trying to time fall seeding, I would recommend that you keep an eye on the 10 day forecast to see when temperatures are starting to cool. But if you get a dry sunny day to plant and it’s reasonably cool, I wouldn’t delay.
So if I want to plant habitat for pollinators and natural enemies next year, what should I do?
First, think about the time, money, and equipment you have available, as well as the area you’d like to plant. There probably isn’t a single right way to establish this habitat, but there may be a best way for you.
You can find more details on the techniques we used (and some links to other resources) here.
This post was written by Amara Dunn, Brian Eshenaur, and Betsy Lamb.
This work is supported by:
Crop Protection and Pest Management -Extension Implementation Program Area grant no. 2017-70006-27142/project accession no. 1014000, from the USDA National Institute of Food and Agriculture.
New York State Department of Agriculture and Markets
In December, I updated you on how perennial wildflowers and grasses were establishing in our beneficial insect habitat plots during the 2019 growing season. As I wrote that post, I quickly realized that there was too much good information for just one post. So here’s the rest of the story when it comes to plant establishment – time, money, and weeds.
