Tuesday, April 15, 2014

Jack Frost Does Not Work Alone


Dead canes of a flower carpet rose
As March was going out like a lamb, a nursery submitted four container-grown shrubs to the PDIC: three rose cultivars and a lilac. Very young shoots on these plants were withering and dying. At least in the case of the lilac – and possibly with the roses, too – the new flush of growth had been hit by the last freezes of the spring. While you’d expect the tender shoots to be blasted by the cold, in this case the woody stems were also dying. Bacterial streaming was seen in much of the stem tissue. We don’t see fire blight on rose or lilac, so what was happening?

The grower suspected Pseudomonas blight. He was right.

A bacterium and its victims

Bacteria were cultured from the stem tissue of the affected plants. Since only Pseudomonas species were of interest, only colonies fluorescent* on a special agar medium were chosen for further work-up. Unfortunately there are a lot of nonpathogenic (non-disease-causing) Pseudomonas species in this world, so it took a little time to sift through the isolates and confirm the diagnosis as Pseudomonas syringae.

Wilting new shoot of a container-grown lilac.
Although Pseudomonas syringae is named after lilac (Syringa), it is capable of causing cankers and dieback in a wide variety of plants. Besides lilac, we’ve found it on the following woody ornamentals: cherry-laurel, flowering quince, Indian hawthorn, Yoshino cherry, and multiple varieties of rose.  In addition, we’ve recovered it from leaf spots of hydrangea and Japanese holly. Bacterial canker caused by Ps. syringae can be a serious problem in peach orchards, but with woody ornamentals we almost always see it in nursery situations. One exception came in last year, on the twig of a weeping willow from a home landscape. As the weather warms up and cankers become inactive, this disease becomes more difficult to detect. According to the PDIC's records, almost every case of Pseudomonas bacterial canker on woody ornamentals since 2008 was diagnosed between February and May. The bacterium is still present on and within plants during the summer, but what I believe is happening is that the hotter temperatures slow down this particular bacterium at the same time as they're (up to a point) invigorating most plants, thus shutting down the disease process temporarily.

Note: We occasionally find Ps. syringae causing leaf spots on ornamentals in the greenhouse, and there are variants – called pathovars – that cause certain very specific problems such as bacterial speck of tomato and angular leaf spot of cucurbits.

How Pseudomonas syringae does its dirty work

Blighted shoots and a Pseudomonas stem canker on rose
Like many bacteria, Pseudomonas syringae is able to live and multiply on plant surfaces. This is known as its epiphytic (“on the plant”) phase. In the recent case, the bacteria were almost surely present before the spring flush occurred, and so were able to strike quickly. These bacteria enter plants following injury, in particular frost damage. What’s more, the bacterial cells actually promote freeze damage through a process known as ice nucleation. How this works is succinctly expressed by Sinclair and Lyon:

"Ice-nucleating strains of P. syringae and certain other bacteria can trigger ice formation in plant tissues cooled to between -2 and -5ºC [28 to 23 ºF] but not acclimated to low temperature. Ice then disrupts cells, causing symptoms of frost damage. In the absence of an ice-nucleating factor, frost-sensitive plants may tolerate brief cooling to these temperatures because water in their tissues remains in liquid form, supercooled." (Diseases of Trees and Shrubs, 2nd Ed. 2005. p.368)

For more information, see this review by Gurian-Sherman and Lindow. You might also check out this laboratory video of ice nucleation by bacteria added to supercooled water.

As if this ice-nucleation trick were not enough, Pseudomonas syringae also produces a toxin that damages plant cells.

How to reduce your losses

The most important way to minimize damage to woody plants from Pseudomonas syringae is to limit the stressors that predispose plants to infection. Stress factors include pruning injury and frost injury. Bacterial canker of stone fruits caused by Pseudomonas syringae can be reduced by pruning in the early summer, instead of the fall or winter. Sanitize shears or knives frequently, and avoid working the plants when wet. Don't overfertilize plants, especially when they need to harden off for the winter. Protect plants during cold snaps. Don't allow plants to undergo stress from too much or too little water. Keep foliage and stems as dry as possible by changing irrigation methods or reducing overhead irrigation, which favors and spreads the bacteria. If you’ve already had this problem, Ps. syringae is probably present as epiphytic populations on the surfaces of much of your nursery stock and even the surrounding weeds. There are few chemical options that hold any promise, at least not enough to make a recommendation.

As I write this, winter is getting ready to take one last shot at North Carolina, with freeze warnings up for the western half of the state. It's another opportunity for Pseudomonas syringae, too.

*Chemist’s Corner:

Colonies of fluorescent pseudomonads photographed under UV light.
The fluorescent pigments of Pseudomonas species are seen by shining a long-wave UV lamp on the cultures. These compounds belong to a class of chemicals called siderophores. If you know Latin, you might think that siderophore means “star bearer”, but in this case the root is the Greek word for “iron”. (A big thanks to Roland Wilbur Brown’s 1956 book Composition of Scientific Words for setting me straight.) Iron is an essential element for microbial growth, and siderophores have the important task of scrounging precious iron from the bacteria’s environment.

Thursday, April 3, 2014

A. destructor

This may look like the toothy mouthparts of some sinister little animal,
but it's really the posterior end of an armored scale (Diaspididae).

Armored scales (Diaspididae) are one of the most common insect pests of ornamentals. More like diseases than insects, these sedentary bugs (literally - they are in the Order Hemiptera) sit and suck the sweet fluids of plants, all the while taking energy from their host and replicating as big sacs of eggs. Their babies, called "crawlers", infest new areas and settle in for the long haul of motherhood (or the short, but free, life of a winged male).

Over the past month or so we received two samples of different plants that had an interesting armored scale infestation. The first was poet's laurel (Danae racemosa), that looked as if it was a variegated variety from the amount of chlorosis associated with the scales:

Poet's laurel (Danae racemosa) leaves with yellow and brown areas due to scale insect pressure. 

Looking under the microscope, I noticed two types of armored scales. The first, and less common, were some typical brown, oyster-shaped fern scales (Pinnaspis aspidistrae). However, the most noticeable scale was one I did not recognize. It had a very thin, translucent test that resembled delicate wax paper with a bright yellow scale underneath:

Although it may look like these scales are under the epidermis of the plant, they are really hiding under a thin test (the covering common to armored scales, to which they owe their name).

When lifted off, the scales underneath looked like this:

Scales with their test removed (top one is facing right and bottom one is facing left).

The scale certainly had the rounded shape characteristic of most members of the subfamily Aspidiotinae (as opposed to the elongate shape of most Diaspidinae, such as the previously mentioned fern scale)...but what was it? Well, I have to confess that our friend and former clinic member Dave Stephan took a peek during a visit and thought it looked like the genus Aspidiotus. Luckily he said that, because the scale book I most often reference first, Scale Insects of Northeastern North America by Michael Kosztarab, does not cover this genus which is primarily Southern in distribution. So I referenced another great resource Ferris's Atlas of the Scale Insects of North America. Lo and behold, Dave was right - the scale keyed out to the genus Aspidiotus and further to the species A. destructor, the coconut scale.

Luckily I had identified that sample, because the same day there was an image of aucuba (Aucuba japonica) that was submitted with a potential scale infestation. Although I was not able to ID the species from the pictures, when the sample came in I recognized the similarities with A. destructor:

The yellow spots on this aucuba leaf are intentional variegation - not chlorosis attributed to the scales. 
Close up of different-sized coconut scales (Aspidiotus destructor).

After clearing some specimens I was able to definitively ID them as the same scale. Was this a coincidence? Probably, but who knows whether these scales are becoming more abundant. Our clinic records show that A. destructor was only submitted and identified four times in the 14 years prior to these two samples. Does that mean that we will be seeing more of this scale? I am not ready to conclude that, but if more are submitted this year we may have to investigate what's going on.

----------

A little more on the scale.

Aspidiotus destructor was described by Signoret in 1869 and goes by several common names including bourbon scale and transparent scale (I am assuming based on the thin test). The scale appears to be Southeast Asian/Pacific in origin, but has been spread throughout the world. Although mainly a pest of coconut and banana, it is extremely polyphagous being found on over 60 families of plants. The genus is characterized by the following traits that can be seen in the title image (Ferris, 1938):

  • absence of paraphyses or intersegmental scleroses
  • three pairs of lobes with no indication of a fourth pair
  • plates long, flat and fringed (two between median lobes, two between median and second lobes, three between second and third lobes, and a variable number beyond third lobe)
  • characteristic sclerotization on dorsum of pygidium

The scale can cause significant economic damage at high densities (which can be common), stunting plants and eventually killing them if enough of the leaves become unable to undergo photosynthesis. Treatments can include chemical control, but there is also a diversity of natural enemies known to attack the species including various fungi, ladybugs, thrips, mites and several parasitoid wasps, most of which are in the family Aphelinidae. In fact, after clearing the scales from the aucuba, I noticed some sinister-looking aliens inside a few of the scales that are certainly wasp larvae and likely a species of Aphelinidae (below "A"):


A cleared female coconut scale showing a parasitoid wasp larva (A), mouthparts/stylets (B), scale egg (C)
and pygidium (D). [Thanks to Mike Munster for helping take pictures of the scale under the microscope]

I don't know whether or not the wasps are able to keep these scales in check alone, but there were at least a few  being eaten by these tiny larvae. Every bit helps I guess!

References:
Ferris, G.F. 1938. Atlas of the scale insects of North America. Series 2. Stanford University Press, Palo Alto, California

Signoret, V. 1869. [Essay on the gall forming insects (Homoptera - Coccidae) - 3rd Part.] Essai sur les cochenilles ou gallinsectes (Homoptères - Coccides), 3e partie. Annales de la Societe entomologique de France (serie 4) 9: 97-104.

Monday, March 24, 2014

Jumpin' Junipers - Red Cedar Problems

Dr. Chuck Hodges at age 82.99.
This blog post is dedicated to our esteemed colleague and expert on tree diseases and on molds, Dr. Charles Hodges, who celebrates his 83rd birthday today. He collected some eastern redcedar (Juniperus virginiana) branches from a golf course last Thursday that demonstrate several problems we'll be seeing over the next few weeks.






One branch was remarkable in that it had not one but two Gymnosporangium rusts, indicated by the arrows in the photograph below.
Infections of quince rust (left) and cedar-apple rust (right) on the same eastern redcedar branch.

The large woody galls on the right are produced by cedar-apple rust (Gymnosporangium juniperi-virginianae), which will be exuding gelatinous orange telial horns when the warm rains arrive in April or May. The spores produced on those horns (basidiospores from teliospores, if you want to get technical) will blow on the wind and infect the leaves and fruit of nearby apple and crabapple trees. Those infections will result in the production of another kind of spore which - if fortunate enough to get a ride on the wind to a juniper - will cause a new infection in summer. Eastern redcedar and Rocky mountain junper are the principal hosts. Those infections will not develop into galls until the year, and they'll mature the following spring. For more information and some nice pictures, see last year's blog post on this disease.



Quince rust infection on an eastern redcedar branch.
The smaller orange-colored swelling on the branch is the telial stage of quince rust. By the last week of March we already see this one forming spores on juniper in eastern North Carolina. The gelatinous telia function the same way as those of cedar-apple rust but are not as large and showy. Another difference is that quince rust infections on juniper are perennial, whereas the cedar-apple gall dies out after producing its spores.
The quince rust fungus, Gymnosporangium clavipes under the microscope.
The two-celled orange teliospores are only 1/500 of an inch long.
Carrot-shaped pedicels beneath are diagnostic for the species.
Quince rust affects not only quince and flowering quince but also hawthorn, serviceberry, and very commonly ornamental pear, where it sporulates abundantly on fruits and less so on swollen twigs in the early summer.
Ornamental pear fruits covered with the white papery peridia of the aecia of quince rust.
Shed spores from the fruit give an orange cast to the leaves. Note: This stage is still months away.
 What about control measures? On juniper you can prune out the galls if they are unsightly or if the branch dies. On susceptible cultivars of apple grown in the vicinity of junipers, fungicide sprays may be needed to avoid losses.



Symptoms of Kabatina tip blight on juniper.
The other disease that Chuck brought in was Kabatina tip blight. The last several inches of the affected twigs had died and faded. When tip blight occurs on juniper in North Carolina in the late winter, Kabatina is the prime suspect. This is also a fungal disease, but you have to look hard with young eyes or a handlens in order to see the tiny gray/black spore-producing bodies (acervuli) at the base of the dead twig. These spores are probably moved by rain splash rather than the wind. In this case the spores are capable of infecting juniper rather than some alternate host. Infection requires some sort of injury either by insects or physical damage. References disagree on whether infection occurs in the fall or spring, but the tip does not die until the following year. Not just eastern redcedar but also other Juniperus spp. are susceptible. No control measures are needed.

Monday, March 17, 2014

These blow flies needed a home...

The color and size range of Lucilia coeruleiviridis is amazing, from blue to green, even bronze. The absence of certain little hairs and other traits unite these guys and gals.

... but not your home. This is about the many dead specimens of blow flies (Calliphoridae) that take up residence in the North Carolina State University Insect Museum.

It all started one day in December when I came across a lovely blow fly resting on the side of the university parking deck. I couldn't help but capture it in the little bag I keep in my wallet - it would make a great subject for photos once I got into the clinic! After settling in and letting the fly calm down under a container on a piece of white paper (a tried and true technique), I snapped some pictures:

Male Chrysomya megacephala have different sized facets on the top and bottom of each eye.

I recognized it as a male Chrysomya megacephala, or the unfortunately-named oriental latrine fly, one of two introduced species of Chrysomya in the United States (the other being C. rufifacies). After some modeling, the fly felt it was time to leave and, despite my attempts to catch it, I lost it in the clinic. Oh well. I love to have the specimen along with photos, but surely our collection was bursting with specimens of this seemingly-common invasive fly? Apparently not.

Looking in the three drawers of blow flies we have in the museum using our online GigaPan images, I found not a single specimen of the fly that just got away! In fact there wasn't even a tray for the genus Chrysomya. This had to be rectified. So my project for the winter was to organize the drawer and a half of unsorted specimens to see how many species we might have, awaiting identification.

- - - - - - - - - -

When the samples started to slow down  in the clinic around January, I walked upstairs to retrieve the drawers of potential treasures. My first step was to find the main literature on identifying blow flies. Luckily, I knew they were a relatively small (84 species in the continental US) and easily identified group, in part because of their importance to humans and animals. Terry Whitworth's key to the genera and species of most of our blow flies served me very well, as did the very nice web key put out by the Canadian Journal of Arthropod Identification. I also knew there were some exotic specimens and luckily a few were covered in the generic key in the Manual of Central American Diptera Vol. 2.

After finding the means to sort through the flies, where would I start? First I got rid of all the obvious non-blow flies, mostly other calyptrates (Tachinidae, Muscidae, Anthomyiidae and Sarcophagidae), but also some shiny Syrphidae.

I didn't want to tackle the two most common blow fly genera (Lucilia and Calliphora; see below). So next I started grabbing out the most obvious: the striped ones! I am talking about screwworm flies (Cochliomyia). The primary screwworm, Cochliomyia hominivorax, is considered to have been eradicated from North America since 1966. It was eradicated because its larvae infest and damage healthy tissue in both animals and man, sometimes leading to death. This is certainly a fly we don't want around. I didn't find any specimens of that species in the unsorted material, but did find many of its common, close relative the secondary screwworm, C. macellaria. These differ greatly from their medically and veterinarily important cousins, in that they only feed on rotting carrion (so unless you're a zombie, there's nothing to be worried about).

A male primary screwworm fly (Cochliomyia hominivorax) tagged for research.
Peggy Greb, USDA Agricultural Research Service, Bugwood.org

At this time I also noted many other specimens with dark faces and dark blue/green bodies, which turned out to be black blow flies (Phormia regina). They made up a large portion (114 specimens) of the material I was looking at, so once they and the Cochliomyia (64 specimens) were sorted out it looked like I was making progress.

Now I felt like Neo from The Matrix: which color pill do I take? In front of me was a sea of large blue flies interspersed with equally as many green or copper flies, all with varying degrees of shine. I decided to go with the smaller green ones that I assumed were all Lucilia, the very common green/bronze blow flies. Most people are familiar with these since they enter homes in search of rotting meat on which to lay their eggs. They are also commonly used in post-mortem forensics (along with other blow flies that feed on remains). Some are even used in maggot therapy - the process where maggots are applied to a wound and feed only on the decayed, bacteria-ridden tissue, effectively cleaning the wound and allowing it to heal properly. If you've seen the epic movie Gladiator you know what I am talking about (and it is still used today).

Green bottle/blow flies (Lucilia) have a blue-green or coppery bronze appearance and are commonly encountered.

Most (174 specimens) of this genus turned out to be Lucilia coeruleiviridis (see title picture), while we also had some L. cuprina (26 specimens), L. sericata (59 specimens) and L. illustris (20 specimens). The most interesting was one specimen of Lucilia silvarum. The species was formerly placed in the now defunct genus Bufolucilia, whose name relates to the fact that females have a habit of laying eggs on toads (genus Bufo) or frogs. The larvae hatch and bore into the amphibian's skin, sometimes in the nose or eyes, and often kill the host. Truly grotesque (for those who are not squeamish, here is what it can look like).

Not all small, metallic green flies were blow flies, however. Among the Lucilia were some impostors, most of which (51 specimens) were a species called Neomyia cornicina. This member of the family Muscidae breeds in cattle dung and, although very similar, has some small differences including a much more metallic face and only one pair of postsutural acrostichal bristle (all blow flies have at least two pairs).

Now onto bigger and bluer flies: Calliphorinae! This group includes the largest blow flies around. Most are dull to shiny blue with some golden or silvery areas on the head or abdomen, and are carrion feeders (though will also come to dung).
Calliphora (like this C. livida) are large, blue blow flies that are often are the loudest flies buzzing inside homes.


I found a number of Cynomyia cadaverina (45 pretty blue-green-purple flies with an ominous name) and a single specimen of Cyanus elongatus, which I was happy to add as a new species to my favorite website Bugguide.net (specimen here). Most of the large blue flies, though, were one of several species of Calliphora, of which the following species (specimens) were identified: Calliphora stelviana (2), Calliphora aldrichia (1), Calliphora latifrons (3), Calliphora vomitoria (12), Calliphora vicina (74), Calliphora terraenovae (9), and Calliphora livida (7).

The last group of blow flies to receive my attention, also happens to be the oddest of the bunch. Cluster flies (Polleniinae: Pollenia sp.) are dull brown or gray with lots of crinkly golden hairs on their thorax - very different from their metallic brethren:
A male cluster fly (Pollenia sp.) resting on a leaf. Note the golden hairs on the thorax, diagnostic for the genus. 

Unlike most other blow flies that mainly feed on carrion or dung, cluster flies are parasites with an unusual host: earthworms! Females lay eggs on the ground where earthworms are present. The young larvae are able to attach to and penetrate the worm to feed on its insides, all the while sticking its back end out of the host to breathe. Sometimes multiple specimens infest a single host, though this can lead to the death of all parties involved. Cluster flies are also known for entering homes in the fall to overwinter, as discussed previously on this blog. Pollenia in the US were traditionally treated as a single species, P. rudis, but more recently have been divided into a several species. I took all the specimens labeled as P. rudis and all the unsorted specimens and found the following three species (specimens): Pollenia rudis (73), P. pediculata (28), and P. angustigena (33).

And what about those Chrysomya? Well, I ended up finding four specimens of the species (C. megacephala) that got away and another two specimens of the other species, C. rufifacies. So at least we have a few, and I will be sure to collect more when I see them.

- - - - - - - - - -

All together I looked at over 800 blow fly specimens and added 11 species previously unidentified for the collection. Many were from NC, but there were also some from the North or out West, and good series from Alaska. We also had one specimen of Hemilucilia segmentaria from Costa Rica, new for our collection. The oldest specimens (but still surprisingly pristine) were collected in the early 1900's, some even by our most famous entomologist Clement S. Brimley (who together with his brother, Herbert H., helped found and grow the NC Museum of Natural Sciences).

I hope to do this for other groups in the museum as it serves many important purposes. Specimens are easier to find for loaning to researchers and we have a better idea of what holdings we have. We can also better organize the little insects on pins to free up room for more specimens. On a personal note, doing these identifications gives me practice with these groups and allows me to see the literature and types of features that are used to identify them.

Maybe I'll tackle the crane flies next...only 15,000 species in the world!

Resources:
Keys to the genera and species of blow flies (Diptera: Calliphoridae) of America north of Mexico (2006) T.L. Whitworth. Proceedings of the Entomological Society of Washington 108: 689–725, 2006http://www.birdblowfly.com/images/Publications/Keys.pdf

Blow flies (Diptera; Calliphoridae) of eastern Canada with a key to Calliphoridae subfamilies and genera of eastern North America, and a key to the eastern Canadian species of Calliphorinae, Luciliinae and Chrysomyiinae (2011) S.A. 
Marshall, T. Whitworth, and L. Roscoe. Canadian Journal of Arthropod Identification No. 11, 11 January 2011, available online at http://www.biology.ualberta.ca/bsc/ejournal/mwr_11/mwr_11.html, doi: 10.3752/cjai.2011.11

Monday, February 10, 2014

Trouble for the Sweet Green of the South

Collard greens are a staple of southern cooking. Slow cooked with the salty flavor of ham hocks and served up with golden corn bread, collards make the perfect meal for a cool winter’s day. Collard greens belong to a group of loose-leaf cultivars (non-heading) of Brassica oleracea. Other members of Brassica oleracea include cabbage, broccoli, Brussels sprouts and cauliflower. These leafy greens are believed to have descended from Asian wild cabbages and eventually made their way into Europe where ancient Greeks and Romans grew the plants in domestic gardens over 2,000 years ago. When colonists settled in the New World, greens were a major part of any well-rounded vegetable garden. In the South, collards were abundant and cheap and quickly became a main ingredient in soul food inspired dishes that southerners still enjoy today.

Lately, we have seen quite a few collard and cabbage samples in the clinic. This season, white mold has been quite prevalent in both home gardens and commercial fields. White mold is caused by the soil-borne fungus Sclerotinia sclerotiorum. This disease is favored by cool, wet conditions and most infections occur in late winter to early spring. Early symptoms of the disease are tan, water-soaked, circular areas on the leaves or crown. These areas quickly become covered in fluffy white fungal growth, followed by a watery breakdown of infected tissue. Eventually, the fungus completely colonizes the head and produces large, black, seed-like structures called sclerotia.
White mold (Photo: E. Lookabaugh)
Sclerotia can survive in the soil for many years. Cool, wet conditions trigger the sclerotia to germinate. Sclerotia can germinate to form mycelium that directly infects plant tissue, or to produce small cup-shaped apothecia (fruiting bodies). These small, mushroom-like apothecia are most likely to form when the sclerotia have been exposed to cold, saturated soil for several weeks. They produce millions of ascospores that can be windblown to nearby plants and infect.
Germinated sclerotia with mushroom-like apothecia (Photo: H.D. Shew)
Sclerotinia sclerotiorum has a very wide host range and can attack over 350 species of plants. Common vegetable hosts include cabbage, broccoli, cauliflower, Brussels sprouts, lettuce, pepper, bean, pea, potato, and tomato. Field crop hosts include sunflower, canola (rapeseed), and soybeans. Additionally, several species of weeds are hosts, including ragweed, dandelion, wild clover, vetch, common chickweed, and pigweed.
Sclerotinia on field grown broccoli (Photo: B. Shew)
Controlling white mold can be difficult and requires an integrated approach. Maintain good air circulation by using proper plant spacing and selecting varieties that have minimal leaf overlap between adjacent plants. Maintain good soil drainage by planting in raised beds. Planting susceptible plants in areas with a history of white mold is not recommended. Rotate with non-susceptible plants such as small grains (rye, wheat), corn, beets, onion, and spinach to prevent inoculum build-up. Only very long rotations (4 years or more) will be helpful in areas with high inoculum levels. Practice good weed control strategies to remove weedy hosts from the area. Planting a small grain as a winter cover crop may help to exclude winter weed hosts. Deep plowing or turning of the soil can be useful because sclerotia typically do not germinate at soil depths greater than 5 cm. This is only effective for one year as subsequent tillage will bring the sclerotia back up to the soil surface. Avoid spreading the fungus with equipment or on infected planting material. When practical, remove infected plants by bagging and burning or disposing of offsite. Do not place infected plants in nearby cullpiles as these can become sources of inoculum. Avoid overhead irrigation and use subsurface irrigation when possible. There are no chemical control options for the home gardener. A few products are available to commercial producers, but correct timing of sprays is critical for successful control. See the Ag Chemical Manual for more information on chemical and cultural control of this disease.

For information on Sclerotinia-caused diseases of other hosts, see our previous blog post.

For information on another disease of collards, see our post on black rot.

Tuesday, January 28, 2014

Winter Insects and a Spring Foreshadowing

With the extreme cold snap and the slow march of winter, most people think that insects and other small animals would be too chilly to make an appearance. Their diminutive size might lead one to think that they would freeze solid out in the environment. For the most part that is true: insects tend to either hibernate or migrate during this time of year to avoid extreme cold. But what about those that enjoy the cold? Many groups only come out this time of year and have special adaptations (such as natural antifreeze - apparently common in Arctic species) to survive the low temperatures. Here are a few cold-loving groups that come to mind:
  • winter crane flies (Trichoceridae) - a family of mosquito-like flies related to their warm weather cousins, the true crane flies (Tipulidae); you may have seen them at your porch light
  • winter stoneflies (Taeniopterygidae) - develop in well-oxygenated streams especially in the mountains; they emerge in the fall, winter and early spring
  • snow crane flies (Tipulidae: Chionea sp.) - these small, wingless, spider-like flies only come out in cold weather, usually being observed on snow
  • snow scorpionflies (Boreidae) - these tiny, hopping insects are only found in the Northern Hemisphere, most commonly in mountainous areas with snow; their larvae feed on mosses
  • snow fleas (Hypogastrura) - often seen in masses on snow, these tiny springtails (not true fleas) enjoy the cold weather
  • some dung beetles (Scarabaeidae) - in my studies on the seasonality of dung beetles in NC, I found that a few species were only present in the cooler months of the year (below)
Aphodius distinctus is one of the dung beetles that is present in the fall and winter, but not during other times of the year (from Bertone 2004).

More types of insects than you think are active in the frigid times of the year*. Most go unnoticed (maybe because we are inside more) and pass their days doing what they do and not bothering us.

Others are a sign of what's to come. Despite their autumnal name, the fall cankerworm moths (Alsophila pometaria) have been out and about for a few months. In fact, just two weeks ago I was alerted to a plethora of females mistaking some campus building pillars for trees. The wingless females were clinging to the cement and some had started to lay their eggs. No doubt others had already done so since they appeared in the late fall.

Female fall cankerworms are strange moths that lack wings (though not unique among Lepidoptera).

The tiny, barrel-shaped eggs of the fall cankerworm laid in a nice batch.

I was also surprised by the abundance of male fall cankerworms at lights and elsewhere this year - something I have not particularly noticed in the past. Males, unlike the wingless females, have large, drab wings and look...rather moth-like and fairly mundane.

Male fall cankerworms are fully-winged and can be found at lights during the cooler months.
So what does this mean? I am assuming the abundance of adults is a direct result of the previous spring's many, many larvae that plagued local trees and bushes, raining down from above on silken strands. But does that mean that we will have as many (or, even more) this coming spring? I am not sure, but we will know in a few months when these little guys come out:

Fall cankerworm caterpillars are all too familiar "inchworms" in recent years.
Oh, and don't forget about spring cankerworms! The silver lining? At least it will be warm outside.

*there is even a whole group of insects, the ice crawlers (Grylloblattodea) that only exist in the cold mountains of the Northern Hemisphere, though in the US only in the Western part of the country; they cannot live much above freezing and will die sitting in a person's hand!

Monday, January 13, 2014

Nematodes Chilling Out Now, But Ready for Action

Nearly dead liriope plants with crown rot and root-knot nematodes.
Last week we received a sample of liriope from NCSU’s Centennial Campus. The main problem was Fusarium crown rot, but it also suffered from root-knot nematodes (Meloidogyne). Based on a microphoto I took, Dr. Weimin Ye of the NCDA&CS Nematode Assay Laboratory gave a tentative identification as the southern root-knot nematode, Meloidogyne incognita.

Root-knot nematodes are microscopic roundworms that are parasitic on cells found in plant roots. The nematodes have a spear-like stylet that allows them to pierce the root cells and feed on the nutrients found there. Root-knot nematodes also use the stylet to inject the cells with nematode "saliva" which contains substances that, among other things, cause the cells near the nematode's head to swell. This swelling eventually leads to the galls that are a symptom of root-knot nematode infection.
  
You may be surprised to know that we’ve found root knot nematode damage on samples from every month of the year, although about 80% of them arrive between mid-May and the end of October. In a blog post last month, Dr. Barbara Shew made mention of this particular pest in the context of the relatively unknown “Nematode Song”.

Meloidogyne species have an extremely wide host range. The PDIC has made a confirmed or suspected diagnosis of root-knot nematodes on many North Carolina host plants since January 2008. This list is by no means exhaustive. For example, figs are a known host that did not show up during this period.

Ajuga, Angelonia, Aucuba, Azalea, Beans, Beets (garden & sugar), Begonia, Bermudagrass, Boxwood, Butterfly-bush, Cockscomb, Coleus, Gardenia, Corn, Cotton, Creeping bentgrass, Cucurbits (Cantaloupe, Cucumber, Watermelon), Euphorbia, Foxglove, Hydrangea, Impatiens, Itea, Japanese holly, Lantana, Liriope, Okra, Pansy, Peach, Peanut, Pentas, Potato, Sasanqua camellia, Soybean, Spinach, Strawberry, Sweetpotato, Tobacco, Tomato.

Plants listed in bold are those with three or more diagnoses during the last six years. The plants most often diagnosed were boxwood (6 samples), cucumber (7), tobacco (5) and tomato (12). Of course this is due in part to the popularity of these plants in North Carolina.

Not all species of Meloidogyne are created equal. For example, all four of the most common species can reproduce on tomato, but only two of them are problems on peanut. Even within species there are races that differ in their host ranges.

Rather than repeat the very useful information on root-knot nematode damage and control from Charlotte Glenn’s previously cited article, I’ll just add a few comments. One is that if your interest is bedding plants, the University of Florida has a publication showing the relative susceptibility of various cultivars.

Charlotte mentioned how to tell a root knot from a legume nodule. Another challenge can be distinguishing between root knots and the normal storage swellings on roots of plants such as daylily, liriope and mondograss. In my experience, the average tuberoid swelling is larger than a root knot, but gall size can vary with the host plant. When in doubt, you can do a dissection under a magnifying glass or a stereomicroscope.
Root-knot galls (left) and swollen storage area (middle) on liriope roots.
The tip of a standard pencil (right) is for size comparison.
Choose swellings that are solid, then shave or pick off layers with a scalpel or razor blade. Look for the pearly white swollen posterior of the adult female nematodes. They are easily “popped”, so work carefully. With a little more effort you can lift out the entire nematode and see the narrowed head and esophageal region.

Two root-knot nematodes dissected out of their gall (arrows).
One is in hind view, the other in side view. Scale bar=0.4mm.
An interesting point to ponder at this time of year is how nematodes survive the “big chill” of January, especially in the absence of a host plant. The short answer is: eggs. Adult females are pretty much egg-producing machines, extruding them in a gelatinous matrix throughout the growing season. Egg masses can be seen on the gall surface if you look under magnification before washing the roots. They may be brown or white.
Brown masses on the surface of these galls are nematode egg masses.

An important part of the nematode’s reproductive “strategy” is that not all the eggs hatch as soon as conditions become favorable; some remain inactive. This inactive period is known as diapause and confers a survival advantage: if conditions improve only temporarily, the unhatched eggs will survive. Fortunately for our plants, root knot eggs will not survive more than about one year if they can’t hatch and infect a susceptible plant root. In this week’s sample, eggs at different stages of development could be seen in the same mass. Some eggs showed no signs of further development, while others already had active juvenile worms inside. In the photo below, you can even see the stylet of the unhatched juvenile. If you have doubts as to whether this nematode was alive, click here.
Meloidogyne eggs within an egg mass. Those at top and left are
undifferentiated. In the egg at right there's a well-developed juvenile.
There are limits, of course. Many root-knot nematode species will not survive the extreme winters of our northern states. That doesn’t mean that they don’t have nematodes up there. My first plant pathology professor, Dr. Dave MacDonald at the University of Minnesota, told us about a state trooper who found live nematodes after melting dirty snow from a blizzard that had picked up soil from points farther west.

There is so much more that can and should be said about these fascinating creatures. Here are two excellent print resources, used in the preparation of this blog. The third is an online introduction to the topic, from the American Phytopathological Society.
  • Shurtleff, M.C., and Averre, C.W. 2000. Diagnosing Plant Diseases Caused by Nematodes. APS Press.
  • Sasser, J.N., and Carter, C.C., eds. 1985. Meloidogyne: An Advanced Treatise. NC State University Graphics.
  •  Mitkowski, N.A. and G.S. Abawi. 2003. Root-knot nematodes. The Plant Health Instructor. DOI:10.1094/PHI-I-2003-0917-01 Revised 2011  http://www.apsnet.org/edcenter/intropp/lessons/Nematodes/Pages/RootknotNematode.aspx
Note: Many other genera of nematodes are important pests of plant roots, although most don’t cause galls. These tend to live in the shadow of their highly destructive root-knotting cousins, and we’ll give them some attention in a future blog.

Mike Munster and Barbara Shew

Friday, December 20, 2013

The Heartbreak of Botrytis

Decay at the terminal portion of the flowering stem of
poinsettia, due to infection by Botrytis cinerea

Dark blotches are bract
infections by B. cinerea
On Monday, December 16th, we received a sample of poinsettias that for all commercial purposes had been ruined by Botrytis blight. Botrytis cinerea is a pernicious and ubiquitous fungus that particularly infects wounded or senescent tissue such as old flowers. From this foothold it can spread to other plant parts. When the fungus sporulates, the colors of the black conidiophores (spore-bearing threads) and white conidia (spores) combine to give the appearance of a gray mold. The spores are easily carried around on air currents.

Botrytis cinerea can wreak havoc with many different host species, even causing canker on rose canes and fruit rot on plants such as strawberry. More specialized species of Botyrtis also exist. One is Botrytis elliptica, which affects primarily lilies. Botrytis tulipae causes a disease called "fire" on - you guessed it - tulip.

Closeup of conidiophores and conidia of Botrytis cinerea,
growing on cyathia (true flowers) of poinsettia. Black bar = 1mm.
The Achilles heel of Botrytis is its need for abundant moisture. In greenhouses, care must be taken to ventilate, even for a while after sunset, in order to keep the relative humidity down. Watching watering practices (timing, drainage) is also important. Fungicides are sometimes needed. Another essential element in Botrytis management is prompt removal of dead plant material from the house. For more information on Botrytis in greenhouses, see our blog from May 18, 2012.

Pansy bed at NCSU in early 2012. Botrytis blight.
Be on the look out for Botrytis blight on pansies in the landscape, especially in the late winter and early spring. To prevent problems, avoid overhead watering if possible, and make sure the plant spacing and surrounding shrubbery allow for good airflow. For more information on Botrytis in the landscape, see the May 31, 2013 blog.
Light colored dead blotches on flowers, spreading to leaves,
are a hint that you might have Botrytis blight in your pansies.


Just a reminder to check our holiday closing schedule. We look forward to seeing you in 2014!

Mike Munster and Kelly Ivors