Figuring out Foxglove Aphid Control

The spring bedding crop season is over, so now’s the time to reflect on what worked, and what didn’t, for foxglove aphid control. This way we can prepare for their re-appearance in the fall.

Foxglove aphid feeding on pansy. Note the two dark-green spots on the abdomen and the dark leg joints which are characteristic of this pest. — Foxglove aphid feeding on pansy. Note the characteristic dark-green blotches on the abdomen, and the dark joints of both the legs and antennae. Photo by S. Jandricic, OMAFRA.

To recap, foxglove aphid (Aulacorthum solani) is a “cool weather pest”. It prefers temperatures between 15-25 C, and can’t survive in the greenhouse in summer ⁽¹⁾. Unlike other aphid pests, foxglove aphid tends to feed in hidden locations – primarily the lowest leaves of plants – making it difficult to detect and treat.

And, unlike green peach and melon aphid, biological control of foxglove aphid is definitely a challenge.

Some growers and consultants have been trying both Aphidius ervi and Aphidius matricariae for foxglove aphid. But, as demonstrated by the Buitenhuis Lab here at Vineland, foxglove aphid is a terrible host for A. matricariae, and this wasp will barely parasitize it (see graph). Further, using A. matricariae for foxglove aphid can actually end up spreading this pest, and it’s damage ^(2). This is because the wasps simply pesters the aphids to the point where they drop of the plant and go find a quieter place to eat. So it’s pretty clear that releasing A. matricariae is simply a waste of money and effort if you’ve got foxglove aphid.

: Parasitism rates of foxglove aphid by different Aphidius species. Wasps were offered 50-60 2nd instar aphids. Tests were done in small plastic containers.

But what about A. ervi? Although parasitism rates were high in the lab (73%), results were not as good in practice. Tested in the greenhouse, A. ervi was able to offer about 50% control of foxglove aphid after 1 release. Repeated releases may offer greater control, but this is still not reassuring when you’re talking about a pest where populations can explode quickly.

Why A. ervi does a great job of parasitizing foxglove aphid the lab, but not the greenhouse, is something we’re currently investigating. But until we have an answer, it seems that pesticides may currently be the best option for control of foxglove aphid in floriculture IPM programs. With the current limitations on neonicotinoids, growers will want to turn to Beleaf or Endeavor. (But, since these two chemicals have similar modes of action, and it usually takes several sprays of either to provide complete control, you may want to consider rotating these chemicals with Enstar II to prevent resistance).

If you’ve had infestations of foxglove aphid in your greenhouse, and have anything to say about it’s control, feel free to leave me a comment!

(1) S.E. Jandricic, S.P. Wraight, K.C. Bennett, and J.P. Sanderson. 2010. Developmental times and life table statistics for the aphid Aulacorthum solani (Hemiptera: Aphididae) at six constant temperatures, with recommendations on the application of temperature-dependent development models. Environmental Entomology 39(5): 1631-1642.

(2) L. M. Henry, J.A. Bannerman, D.R. Gillespie2, and B.D. Roitberg. 2010. Predator identity and the nature and strength of food web interactions. Journal of Animal Ecology doi: 10.1111/j.1365-2656.2010.01723.x.

Banishing Broad Mite – New post in Floriculture IPM Blog

I’ve been getting a lot of calls lately about Broad Mites (Polyphagotarsonemus latus) in crops like New Guinea impatiens, torenia, begonias, exacum, ipomea and gerbera. Broad mite can also attack chrysanthemums, so it’s time to start thinking about control of this pest as you’re sticking your new cuttings. Read on for tips on monitoring and control.

Continue reading “Banishing Broad Mite – New post in Floriculture IPM Blog” →

A Crime Against Callies: Floriculture IPM of Black Root Rot

Diagnosing a plant with vague symptoms like wilting, yellow and stunting is much like being a “plant detective”. First, you need to profile the “victim” — here, Callibrachoa plugs. Then, collect DNA evidence. Finally, use knowledge and instinct to narrow down your “suspects”. Only then can you come up with a plan to stop the assailant.

Yellowed plant growth (yellow circle) and dead plugs (orange circle) on a plug tray of Callibrachoa.

An unhealthy plug; few white roots are visible.

In this case, our DNA evidence (c/o UofG Lab Services) gave me 3 possible suspects: Pythium disotocum, Fusarium oxysporum and Thielaviopsis basicola. But which of these was the real culprit?

P. disotocum is rarely documented as an aggressive pathogen in flowers, so we can eliminate that. Similarly, Fusarium is often “around” at low levels without causing a problem. But, T. basicola, better known as Black Root Rot, is a common problem in Callibrachoa, specifically. Rotting roots are not always directly evident (even under a microscope), but severe discoloration of the foliage is a good clue, since this pathogen produces toxins that result in yellowing. Left unchecked, Black Root Rot causes severe stunting and plant death.

Luckily, there are treatments for Black Root Rot, and preventative measures that can be taken.

A fungus gnat (left) and shorefly (right) caught on a yellow sticky card. Both of these pests can transmit Black Root Rot and other pathogens.

Drenches of Senator 70 WP (thiophanate-methyl) are the best option for dealing with an existing outbreak of Black Root Rot
Black Root Rot is often transmitted by fungus gnats and shoreflies. Management of these insects is key to prevention. They can be controlled with soil applications of nematodes, predatory mites (e.g. Hypoaspis), or insect growth regulars like Dimilin (diflubenzuron). These products are compatible with biocontrol programs for other pests, like thrips and aphids.
A high soil pH encourages the growth of Black Root Rot. pH should be kept below 5.6.
Bio-fungicides containing the beneficial organism Trichoderma harzianum (e.g Rootshield) can help protect plants from this Black Root Rot if applied at planting.

For additional information on Black Root Rot, check out http://plantclinic.cornell.edu/factsheets/blackrootrot.pdf

When “harassment” is a good thing in your greenhouse

Well, now that I’ve gotten your attention, let’s be clear that I’m talking about bugs, here, people. Specifically, Western flower thrips and predatory mites such as N. cucumeris and A. swirskii.

L2 western flower thrips next to a predatory mite egg (left) and 2 adult mites (right). — Second instar (L2) western flower thrips next to a predatory mite egg (left) and 2 adult mites (right). Photo credit: Sarah Jandricic.

We all know that predatory mites only kill and eat the small, first larval stage of thrips (referred to as L1’s). Larger life stages (i.e. L2’s and adult thrips) are simply too large for the mites to kill.

But, that doesn’t mean they don’t help control them.

Predatory mites will repeatedly attempt to kill L2 thrips – attacking them up to 40 times an hour ⁽¹⁾. The L2 thrips are able to fend off these attacks by either running away, or “slapping” mites in the face with their abdomens. You can watch the hilarity that ensues when mites attempt to attack too-large thrips larvae here.

But this “harassment” by mites takes it’s toll on thrips. Because thrips spend more time fending off mites, they spend 30% less time feeding ⁽²⁾. Over time, this translates to 40% less damage on plants with predatory mites, compared to no mites ⁽²⁾. And, the presence of mites can reduce survival of L2 thrips by up to 78% ⁽¹⁾, probably because eating less means the thrips lack the nutritional reserves to complete development.

And all of this is accomplished just through “intimidation” of thrips by mites – not through consumption. Scientists term these “non-consumptive” effects, and we are just starting to learn the importance of these effects in biological control. Research is now suggesting that non-consumptive effects like “harassment” may actually account for 50% of the pest control we see in greenhouses ⁽³⁾.

This makes sense with our thrips example. Not only do mites reduce the feeding and survival of larval thrips, but ongoing research from Cornell University shows that the presence of mites reduces the number of eggs laid by adult thrips, and shortens adult thrips lifespan ⁽⁴⁾. Even the presence of predatory mite eggs on a plant has been shown to “scare” L1 thrips into eating less, according to research from Austria ⁽⁵⁾.

And, it would make sense that the more mites you have, the higher the number of “scary” encounters thrips will have with them, improving control.

So, lets give a hand to the hard working, harassing, predatory mite, who’s doing more than we ever thought in floriculture IPM. Stay creepy, little guys. Stay creepy.

References: (1) Jandricic, S.E., Schmidt, D., Bryant, G., and Frank, S.P, NC State University. Unpublished data. (2) Jandricic, S.E. and Frank, S.P. 2014. Too scared to eat: non-consumptive effects of predatory mites. IOBC/wprs Bulletin 102: 111-115; (3) Preisser, E. L, Bolnick, D. I., & Benard, M. F. 2005: Scared to death? The effects of intimidation and consumption in predator-prey interactions. Ecology 86: 501-509. (4) Loughner, R., and Nyrop, J. Cornell University. Unpublished data. (5) Walzer, A., & Schausberger, P. 2009: Non-consumptive effects of predator mites on thrips and its host plant. Oikos 118: 934-940.