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Identify black insect on Granadilla plant in South Africa

Identify black insect on Granadilla plant in South Africa


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Planted a granadilla tree 3 weeks ago and these guys appeared on the new growth.

Never seen them before.


Identify black insect on Granadilla plant in South Africa - Biology

Some of these can be very confusing at times and if I am not careful, it would be very easy to give a mistaken identity to some critters.
If you look at the beetle above and below this, you can see how similar the markings are and yet they are two different beetles.

The third one below also has an almost identicle patterns but at least the rest if him is lighter in color and is easier to tell apart.

The next two can be quite similar to the untrained eye.

yet once again, they are two totally different beetles.

3 comments:

I'm trying to figure out what that third beetle is called and what it scientific name is, besides Christmas beetle, of course. That's what everyone in SA calls them, but it definitely can't be their real name.

Hi Im looking for the name of a black beetle with round white spots. I found several in my vegetable garden and I would like to know if it is damaging my plants?

Hi there. I was wonderig if you could helpme identify a very small beetle. It is about 4mm in length. I have a picture if I can send it to you. There are literally 1000s on my friend's property and really feeling like a pest.


A-Z list of horticultural insect pests

Read about the Avocado leaf roller pest and the damage it does to avocados, custard apples, coffee, tea and other crops in North Queensland and learn how to control it.

Banana aphid

Information on the Banana aphid insect pest which can spread banana bunchy top virus (BBTV) including description, distribution, hosts, damage and controls.

Banana flower thrips

Information on the Banana flower thrip insect pest, which is found throughout the banana-growing districts.

Banana fruit caterpillar

Banana fruit caterpillar (Tiracola plagiata), insect pest of banana and citrus, description of adult, life cycle, hosts, damage, control options, distribution.

Banana rust thrips

Information on the Banana rust thrip. Bananas are the main host, although the Banana rust thrip has been found in citrus and in some native plants.

Banana scab moth

Banana scab moth (Nacoleia octasema) insect pest of bananas.Description, life cycle, host range, damage and control options.

Banana spider mite

Banana spider mite (Tetranychus lambi) the most important and widespread mite pest of bananas. Description, life cycle, host range, damage and control options.

Banana weevil borer

Banana weevil borer (Cosmopolites sordidus), insect pest of genus Musa inclu. banana (M. sapientum) and abaca (M. textilis).Description, damage,control methods.

Banana-silvering thrips

Information on the Banana-silvering thrip insect pest, which affects bananas, chokos, passionfruit and a number of weed species.

Banana-spotting bug

Banana-spotting bug (Amblypelta lutescens lutescens) insect pest of wide range horticultural crops. Decription, life cycle, damage and control options.

Bean blossom thrips

Information on Bean blossom thrips, which affect French beans.

Black scale

Black scale (Saissetia oleae) insect pest of olives, citrus and gardenia, distributed throughout Queensland. Description, life cycle, damage and control options.

Cicada

Cicada (Macrotristria dorsalis) insect pest, including description, life cycle, distribution, damage, host range, and control measures.

Citrus mealybug

Citrus mealybug (Planococcus citri) is a major and frequent pest of many fruit and ornamentals throughout Australia. Description, life cycle, damage and control.

Clearwing borer

independently. Description Clearwing borer (Carmenta chrysophanes) insect pest in all lychee and longan growing areas in Queensland. Description, life cycle, host range, damage, control.

Corn earworm and native budworm

Information and control options for the corn earworm and native budworm insect pests.

Cryptic mealybug

Cryptic mealybugs pose a significant threat to a range of Australian horticultural industries. Description, life cycle, damage and control.

Diamondback moth

Diamondback moth insect pest information and control options.

Eggfruit caterpillar

Information on the Eggfruit caterpillar. Eggplant is the main commercial host but it also occasionally attacks tomato, capsicum and pepino.

Elephant beetle

Elephant beetle (Xylotrupes gideon) is a pest of pineapple, longan and lychee. Exclude beetles with net or remove manually. There is no chemical control.

False spider mite

False spider mite (Brevipalpus sp.) is minor pest of a variety of fruit crops.Predatory mites control numbers but if fruit loss is high, consider treatment.

Flower eating caterpillar

Flower eating caterpillar information and control options.

Fruit piercing moth

Fruit piercing moth information and control options.

Fruit-spotting bug

Fruit-spotting bug information and control options.

Giant grasshopper

Giant grasshopper (Valanga irregularis) damages a variety of plants inclu. coffee and citrus. Chemical control may be used to spot spray heavily infested areas.

Green vegetable bug

Species fact sheet - Green vegetable bug, Nezara viridula.

Helopeltis

Information on the Helopeltis pest in Queensland, including impacts, distribution and control methods.

Loopers

Loopers insect pest information, including description distribution, damage, and control options.

Lychee erinose mite

Lychee erinose mite (Aceria litchii) is a serious pest of lychee foliage, flowers and fruit.Control can be achieved with a strict program of suitable miticides.

Macadamia nutborer

Macadamia nutborer (Cryptophlebia ombrodelta) damages lychee, logan, macadamia and many ornamentals. Control many be manages by biological and chemical methods.

Mango seed weevil

Mango seed weevil (Sternochetus mangiferae), a minor pest with no serious economic damage to fruit. Control programes use quarantine, hygiene and chemical spray.

Mango shoot caterpillar

Mango shoot caterpillar (Penicillaria jocosatrix) is a minor and frequent pest of mango and cashew in late summer. Control should coincide with growth flushes.

Mound forming termites

Mound forming termites (Nasutitermies spp) can affect cashew, mango and citrus grown in drier areas but are usually not a problem in well-managed crops.

Orange fruit borer

Orange fruit borer (Isotenes miserana) is a minor and sporadic pest of a variety of fruit crops in coastal Queensland.Use chemical sprays when numbers indicate.

Oriental scale

Oriental scale (Aonidiella orientalis) is a minor pest of a wide range of crops and ornamentals. Infestations are best treated with biological control agents.

Passionvine bug

Passionvine bug (Leptoglossus australis) is a minor but frequent pest of many fruit crops crops and some cucurbits. No chemcial control is currently available.

Pink wax scale

Pink wax scale (Ceroplastes rubens) is a minor and frequent pest of many tropical fruit crops but is well managed with a range of biological controls.

Potato moth

Potato moth (Phthorimaea operculella) is a serious pest of tomatoes and is usually supressed by parasitoids. Avoid planting adjacent to susceptible crops.

Queensland fruit fly

Queensland fruit fly (Bactrocera tryoni) is a major pest of many fruits in eastern Australia. Chemcial sprays, parasitoids and orchard hygiene may be used.

Red scale

Red scale (Aonidiella aurantii) infests a range of hosts including citrus, passionfruit. Biological control is crucial so spray only if infestation is high.

Red-shouldered leaf beetle

Information on the Red-shouldered leaf beetle insect pest.

Root-knot nematode

Root-knot nematode, Meloidogyne spp, is a problem in many crops especially tomatoes. Plant resistant varieties and use crop rotation, nematicides for control.

Silverleaf whitefly

Information on the Silverleaf whitefly insect pest.

Soft brown scale

Information on the Silverleaf whitefly insect pest.

Stem girdlers

Stem girdlers are ring-barking weevils involving at least three weevil species causing sporadic damage in all lychee/longan districts. There is no control.

Sugarcane bud moth

Sugarcane bud moth (Opogona glycyphaga) affects bananas with crops planted close to sugarcane suffering more. Dust with chemical when applying bunch cover.

Swarming leaf beetles

Information on the Swarming leaf beetle insect pest.

Tomato russet mite

Tomato russet mite (Aculops lycopersici) affect tomato, chilli and capsicum. Maintain good farm hygiene and destroy old crops and weeds, spray when appropriate.

Two-spotted mite

Two-spotted mite (Tetranychus urticae) has a wide host range and can be a sign of excessive insecticide use. Manage with predatory mites and limited spraying.


Scale Insect Pests

The California red scale is an armored scale with a hard, reddish-brown covering over the adults. It can infest all parts of the plants.
Dennis Navea, ControlBest, Bugwood.org

California red scale (Aonidiella aurantii) is an armored scale pest of citrus and difficult to control with insecticides. The adult female scale infests the fruit, stems, and leaves, and appears as ⅟10-inch, reddish-brown spots or scabs on the plant. The female gives birth to 100 to 150 yellow-colored immatures, called crawlers, which disperse by crawling to find a place to settle down and suck nutrients from the plant parts. These crawlers form hard coverings over their bodies and become immobile adults.

Severe infestations cause leaf yellowing and drop, dieback of twigs and limbs, and occasionally death of the tree. Citrus tree damage is most likely to occur in late summer and early fall when populations of this scale are highest, and moisture stress on the tree is greatest. Naturally occurring parasitic insects may help control some of the scales in outdoor settings.

Citrus snow scale infests limbs and twigs initially, but with a severe infestation, this scale will colonize the foliage.
Central Science Laboratory, Harpenden, British Crown, Bugwood.org

Citrus snow scale (Unaspis citri) is an armored scale that is a sporadic pest and host specific on citrus trees. Heavy infestations can almost completely cover the bark and larger limbs and give a white, snowy appearance. The inconspicuous, immobile female scales are brownish-purple, oyster-shell shaped, and ⅟16– to ⅟11-inch long pests. The snow-white, winged males give the descriptive name to this scale species. Crawlers (immatures) are very small, light orange to reddish and easily spread to other plants and additional branches. Some or all life stages of the scale are found throughout the year (eggs, crawlers, nymphs, and adults). There are multiple generations of this scale during the growing season.

With infestation, citrus tree will have decreased vigor, reduced fruit production, and partial defoliation. Heavy infestations can cause limb and branch dieback, large cracks to form in the bark, and can eventually lead to the death of the tree. Typically, leaves and fruit are not infested until scale populations become severe. Natural parasitoids are unable to keep this scale pest under control.

Florida red scale initially infests citrus fruit and is one of the most damaging scale pests of citrus.
Pedro Torrent Chocarro, Bugwood.org

Florida red scale (Chrysomphalus aonidum) is an armored scale with circular armor made up of three concentric rings. They are dark reddish-brown, have a conspicuous, light brown center, and the size is about ⅟12-inch in diameter. This scale occurs on a wide range of hosts, such as citrus, Aspidistra, and Dracaena, and like most armored scales does not produce honeydew (the sugary waste product that drips from the insects). There may be several generations per year. The immatures (crawlers) are bright lemon yellow. They infest fruit first, and then in late summer and early fall, they feed on the foliage.

Symptoms consist of yellow spots on both the leaves and fruit. If only a few leaves are infested with scales, trim off and dispose of the infested foliage. If the citrus is a landscape plant, sprays are needed to stop the spread of this pest. Heavy infestation may cause severe defoliation. This scale pest is one of the most serious pests of citrus. There are several wasp parasites, which aid in the control of this scale. Additionally, ladybird beetles will feed on the scale crawlers.

Purple scale is also a serious pest of citrus and can infest all parts of the citrus trees. D.R. Miller, US National Collection of Scale Insects Photographs, USDA ARS, Bugwood.org

Purple scale (Lepidosaphes beckii) is an armored scale pest primarily of citrus trees. The adult female scales are small, elongate, ⅟12– to ⅟8-inch long, purple to dark brown, and slightly curved. The adult male scales are smaller. The mobile immatures are very small and white, and there may be two generations per year. This insect pest prefers the shadier and more protected areas of the tree, so the higher populations may be found toward the center of the tree. The dense canopy of foliage protects them from parasites.

This scale infests the citrus foliage, fruit, and stems, and can cause leaf yellowing and drop, spotting and deformity of fruit, shoot malformation, and with heavy infestations, plant death can occur. Symptoms include green spots on fruit that do not color correctly and yellow spots on foliage. A heavy infestation may cause defoliation. Parasitic wasps keep purple scale in check in Florida, but these biological controls may not yet be present in South Carolina.

Florida wax scale (Ceroplases floridensis) are small brownish-purple insects that are covered with a dirty-white waxy covering. They are soft scales that commonly infest citrus, hemlock, azalea, blueberry, camellia, Chinese elm, fig, Chinese holly, yaupon holly, jasmine, mulberry, pear, persimmon, plum quince and other plants. Crawlers (the immatures) are typically pink and are present during late spring or early summer. They migrate to and feed on the underside of foliage for about a month, at which time the female crawlers move to twigs and small branches to continue feeding.

Florida wax scale is a soft scale of citrus and many ornamentals. This scale produces honeydew, which drips onto surrounding foliage. The honeydew is colonized by dark-colored sooty mold and results in foliage becoming blackened.
Chazz Hesselein, Alabama Cooperative Extension System, Bugwood.org

When a scale infestation is heavy, black sooty mold can grow on the clear, sweet, sticky honeydew (the sugary waste product resulting from scale feeding on plant sap) that drips onto nearby foliage. Severe infestations may kill branches. If there is no noticeable blackening of the leaves from sooty mold, then the wax scale infestation is probably not severe enough to kill branches. Typically, natural enemies, such as parasitic wasps, keep Florida wax scale under control.

Cottony cushion scale (Icerya purchase) can be more than a nuisance on shrubs and trees. Host plants include citrus, apple, Nandina, Boston ivy, boxwood, cypress, hackberry, locust, maple, oaks, peaches and plums, pecan, pears, pine, Pittosporum, pomegranate, quince, rose, Verbena, walnut, willow, and other woody ornamentals.

Adult female scale insects have reddish-brown bodies with black legs and antennae. However, the most distinguishing characteristic of this scale is the large, elongated and grooved, cottony-white egg sac. The egg sac (⅜- to ⅝-inch in length) becomes two to 2½ times as long as the body of the female, and there may be hundreds of eggs in each egg sac. Eggs in the egg sac hatch into the six-legged “crawler” stage, which are bright red with black legs. They then move onto larger twigs and branches, feed, and develop through several stages before becoming adults. Populations may increase very quickly during the dryer months of summer.

Cottony cushion scale is another soft scale of citrus. Behind the female scale is a grooved, white egg sac containing hundreds of scale eggs.
Sonya Broughton, Dept. of Agriculture & Food, Western Australia, Bugwood.org

Cottony cushion scale infestations can generally cause older trees to have reduced vigor, premature leaf drop, or twig death, but younger trees can be severely stunted or killed. Similar to the Florida wax scale, these soft scale insects debilitate plants by sucking out sap (phloem), and then excrete honeydew, which coats infested plants. Dark fungi called sooty molds grow in the honeydew. Heavily infested trees become chlorotic and darkened by the mold. During periods of stress, leaves and fruit may drop prematurely, and plants may die.

Chemical Control of Scales: The adult female scales are difficult to control with regular contact insecticides because of their hard, waxy covering. However, sprays of horticultural oil, an excellent, proven product for scale control, kill all stages of scales insects that are present at the time of application. Horticultural oil is safe to use and is an especially good choice for sensitive areas, such as where people are present soon after treatment. Due to its short residual, oil sprays help to conserve beneficial insect species. Horticultural oil sprays control both armored and soft scales.

Apply a horticultural oil spray before new growth begins in late winter or early spring and when the temperatures are above 45 °F. These oils work by smothering overwintering adult female scales, immatures (crawlers), and their eggs. They offer the best control when applied during this dormant season. Oil sprays kill by suffocation. Spray the trunk and limbs with 2% horticultural oil solution to the point of run-off. Make a 2% solution by mixing 5 tablespoons of horticultural oil per gallon of water.

Horticultural oil sprays can be applied any time to control scales any time the temperatures are between 45 and 85 ºF. If scale problems are severe, spring and fall applications may be needed. Additional spray applications may be required when new leaves start to expand in the spring. Make two or more spring applications as necessary at three- to four-week intervals. These springtime sprays provide control of the immatures (crawlers) that hatch after new foliage appears. Spray the trees thoroughly until the oil spray drips or “runs off” from the upper and lower surfaces of leaves, twigs, branches, and the trunk.

When necessary, a 1 or 2% mixture of horticultural oil can be applied again to the foliage during the growing season. For tender new growth, apply a 1% mixture spray (2½ tablespoons per gallon of water). On mature foliage, apply a 2% mixture spray (5 tablespoons of oil per gallon of water). Do not spray in direct sunlight or if rainfall is expected within 24 hours. To lessen the chance of foliar injury and slow the drying time of the oil sprays, apply horticultural oils late in the day.

If citrus fruit are present on the plants, apply horticultural oil no stronger than 3 tablespoons per gallon of water. Shake the sprayer often to keep the oil and water mixed. Examples of horticultural oil products are in Table 1.

Canola oil sprays labeled for horticultural use can also be used to reduce the number of scale insect pests by suffocating all growth stages like the horticultural oil sprays do. Examples of available brands are in Table 1.

In general, using least toxic insecticides, like horticultural oil and canola oil, will prevent harm to beneficial insects. When general contact insecticides are used, they will kill the naturally occurring, beneficial enemies of scale insects. The general contact insecticides will only kill the crawler stage (immatures) of the scale insect because they cannot penetrate the hard waxy covering over the adults. Therefore, contact insecticides should only be applied during the growing season when scale crawlers are present. General contact insecticides registered for insect control on citrus include malathion. For examples of brands containing malathion, please see Table 1.


2. Giant Leopard Moth (Hypercompe scribonia)

This is a large, black, spiny caterpillar. The spines are very sharp and stiff, almost like needles. When it feels threatened, the caterpillar rolls itself up into a ball with the spines sticking out. It also shows bright crimson bands between its body segments. Red and black are universal warning colors, and predators might well think that they&aposre dealing with a wasp. If they do try to take a bite, all they will get is a mouthful of spines.

These beautiful moths, which are large and strikingly marked with black circles and iridescent blue spots, overwinter as full-grown caterpillars, usually under rocks, where you may find them curled up in the middle of winter. In the spring, they spin a cocoon from which the adults emerge in the summer.

The Basics

  • Does it sting? No, but the spines are very stiff and sharp.
  • What does it eat? Plantains, dandelions and violets.
  • Will it seriously damage plants or trees? No. It will eat some of the greens and leaves, but not very much.
  • Is it rare? No, but they are more common in the South.
  • What does it turn into? A really gorgeous moth.
  • Can you raise it to an adult? No, due to its overwintering habits.

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Abstract

It is thought that speciation in phytophagous insects is often due to colonization of novel host plants, because radiations of plant and insect lineages are typically asynchronous. Recent phylogenetic comparisons have supported this model of diversification for both insect herbivores and specialized pollinators. An exceptional case where contemporaneous plant–insect diversification might be expected is the obligate mutualism between fig trees (Ficus species, Moraceae) and their pollinating wasps (Agaonidae, Hymenoptera). The ubiquity and ecological significance of this mutualism in tropical and subtropical ecosystems has long intrigued biologists, but the systematic challenge posed by >750 interacting species pairs has hindered progress toward understanding its evolutionary history. In particular, taxon sampling and analytical tools have been insufficient for large-scale cophylogenetic analyses. Here, we sampled nearly 200 interacting pairs of fig and wasp species from across the globe. Two supermatrices were assembled: on an average, wasps had sequences from 77% of 6 genes (5.6 kb), figs had sequences from 60% of 5 genes (5.5 kb), and overall 850 new DNA sequences were generated for this study. We also developed a new analytical tool, Jane 2, for event-based phylogenetic reconciliation analysis of very large data sets. Separate Bayesian phylogenetic analyses for figs and fig wasps under relaxed molecular clock assumptions indicate Cretaceous diversification of crown groups and contemporaneous divergence for nearly half of all fig and pollinator lineages. Event-based cophylogenetic analyses further support the codiversification hypothesis. Biogeographic analyses indicate that the present-day distribution of fig and pollinator lineages is consistent with a Eurasian origin and subsequent dispersal, rather than with Gondwanan vicariance. Overall, our findings indicate that the fig-pollinator mutualism represents an extreme case among plant–insect interactions of coordinated dispersal and long-term codiversification. [Biogeography coevolution cospeciation host switching long-branch attraction phylogeny.]

Processes affecting the diversification of insects are crucial to understanding the origin of biodiversity, because most animals are either insect herbivores, or natural enemies (predators or parasitoids) of these phytophages ( Novotny et al. 2002). As primary consumers, most insect herbivores are involved in antagonistic interactions with plants and, although herbivores often exhibit host-specific coevolutionary adaptations to plant defenses ( Ehrlich and Raven 1964), recent empirical studies have suggested that host plant lineages are generally older than their associated herbivores ( Percy et al. 2004 Tilmon 2008 McKenna et al. 2009). Such patterns of asynchronous plant–insect diversification are consistent with the general paradigm that insect speciation results from colonization of novel host plants and subsequent reproductive isolation ( Percy et al. 2004 Tilmon 2008 McKenna et al. 2009 Fordyce 2010).

Phytophagous insects are often enemies of plants, but some engage in beneficial pollination mutualisms. A charismatic example involves the ca. 750 species of figs (Ficus, Moraceae) and their pollinating wasps (Hymenoptera, Chalcidoidea, Agaonidae) ( Fig. 1). Agaonids are the only pollen vectors for fig trees and agaonid larvae feed exclusively on the flowers of their Ficus hosts. Each partner is thus entirely dependent on the other for reproduction. Figs are also a major resource for frugivores and most animal-dispersed tropical tree species interact with vertebrates that also consume figs ( Howe and Smallwood 1982). The fig-pollinator mutualism is therefore ecologically important in most tropical ecosystems ( Shanahan et al. 2001). Many fig species reproduce irregularly, are relatively inaccessible in the forest canopy, or today are found only in rainforest remnants, such that coordinated sampling of Ficus and pollinator species for systematic study is difficult. These sampling challenges, coupled with the limitations of analytical tools for large data sets, have hindered progress toward understanding the global evolutionary history of the mutualism, despite the fact that many details of this intricate symbiosis were described almost a century ago ( Janzen 1979 Wiebes 1979 Weiblen 2002 Cook and Rasplus 2003 Herre et al. 2008).

Classification and worldwide distribution of Ficus. The numbers of species per subgenus is represented as a proportion of total Ficus species richness. Breeding systems are indicated as either monoecious (M) or dioecious (D) and modes of pollination are indicated as passive (P) or active (A). *Agaon, Alfonsiella, Allotriozoon, Courtella, Elisabethiella, Nigeriella and Paragaon. **Deilagaon and Waterstoniella.

Classification and worldwide distribution of Ficus. The numbers of species per subgenus is represented as a proportion of total Ficus species richness. Breeding systems are indicated as either monoecious (M) or dioecious (D) and modes of pollination are indicated as passive (P) or active (A). *Agaon, Alfonsiella, Allotriozoon, Courtella, Elisabethiella, Nigeriella and Paragaon. **Deilagaon and Waterstoniella.

Species-specificity in fig pollination appears to be extreme compared with most other insect pollination mutualisms. Most fig species are pollinated by only one or a few wasp species and most wasps are associated with just a single fig species ( Cook and Rasplus 2003 Molbo et al. 2003 Cook and Segar 2010). Pollinators are specifically attracted to volatile compounds emitted by figs ( Hossaert-McKey et al. 1994) and access to the specially modified inflorescences is by means of distinctive mandibular appendages and detachable antennae ( van Noort and Compton 1996). Pollination is either active (two-thirds of the fig species) or passive (one-third, mostly within subgenera Pharmacosycea, Ficus, Synoecia, and Urostigma) ( Kjellberg et al. 2001). Active agaonid wasps collect pollen from the anthers of their native figs and store it in thoracic pollen pockets ( Galil and Eisikowitch 1968 Ramirez 1978). Once inside a receptive fig, they remove pollen from their pockets and deposit it on the flower stigma each time they lay an egg ( Galil and Eisikowitch 1968 Kjellberg et al. 2001). Passively pollinated figs produce large quantities of pollen through anther dehiscence and wasps are covered with pollen ( Galil and Neeman 1977) before flying away from their natal figs.

Closely matching fig and pollinator traits might be products of coadaptation ( Ramirez 1974 Wiebes 1979, , 1982a Kjellberg et al. 2001 Weiblen 2004) but, regardless, trait-mediated interactions have the potential to simultaneously affect the evolution of reproductive isolation among pollinator and fig populations this is because fig wasps breed exclusively in pollinated figs. This line of reasoning has underpinned the hypothesis that cospeciation might account for patterns of fig and pollinator diversity. However, this notion runs contrary to the paradigm that insect speciation generally involves host-switching ( Tilmon 2008) and so it remains a controversial proposition that requires rigorous testing.

Under the cospeciation scenario, phylogenies of figs and pollinators are expected to show substantial congruence. There is some evidence for this pattern ( Herre et al. 1996 Machado et al. 2005 Rønsted et al. 2005 Cook and Segar 2010 Cruaud et al. 2011a), but recent studies have countered the underlying case for cospeciation with evidence of cryptic wasp species and relaxed partner specificity. At least 50 fig species are now known to have multiple pollinator species ( Michaloud et al. 1985, 1996 Rasplus 1996 Kerdelhué et al. 1997 Lopez-Vaamonde et al. 2002 Greeff et al. 2003 Molbo et al. 2003 Haine et al. 2006 Moe and Weiblen 2010 Chen et al. 2012) and as many as 4 different wasp species are known to pollinate a single fig species ( Machado et al. 2005 Cook and Segar 2010). Such cases occur in a broad taxonomic and geographic spectrum, although cases of pollinator species sharing multiple fig species have been reported mostly from monoecious figs in the Neotropics ( Molbo et al. 2003) and the Afrotropics ( Erasmus et al. 2007 Cornille et al. 2012 McLeish and van Noort 2012). In any event, evidence of relaxed host specificity and some incongruent fig-pollinator phylogenies ( Machado et al. 2005) suggest that host shifting is a viable alternative explanation for fig-pollinator diversification.

Cospeciation has been hypothesized for the vertically transmitted endosymbionts of insects (e.g., Moran 2001 Jousselin et al. 2009) but this is not a plausible general model for the evolution of plant–insect associations, which are horizontally transmitted and not so integrated metabolically. Further, if the plant traits that mediate insect associations happen to be phylogenetically conserved, then host shifting among close relatives could also result in topologically congruent phylogenies ( Percy et al. 2004). In addition, historical biogeography has the potential to confound the explanation of such patterns if synchronous plant–insect dispersal to new environments, followed by geographic isolation, results in cospeciation.

Another useful approach is to investigate patterns of temporal congruence ( Page and Charleston 1998). Divergence time estimates for fig and pollinator clades are expected to be approximately equal in the event of coradiation, whereas insect lineages are expected to be younger than hosts in the case of host shifting ( Percy et al. 2004 Tilmon 2008 McKenna et al. 2009).

Previous comparisons of fig and pollinator phylogeny have yielded rather different insights on the relative importance of host shifting and codiversification depending on the taxonomic scope of sampling ( Cook and Segar 2010). Molecular phylogenetic trees appear roughly parallel when based on exemplars of Ficus sections and wasp genera ( Herre et al. 1996 Jackson 2004 Cruaud et al. 2011a), but such deep taxonomic sampling is unlikely to detect host shifts among close relatives ( Machado et al. 2005). On the other hand, regional comparisons of particular fig and pollinator clades have tended to reject cospeciation in favor of host-switching ( Machado et al. 2005 Marussich and Machado 2007 Jackson et al. 2008 Jousselin et al. 2008), although not always ( Weiblen and Bush 2002 Silvieus et al. 2008). A global test for codiversification therefore requires dense sampling of many fig and pollinator lineages across the entire geographic range, but a problem of this magnitude poses a further methodological challenge.

Tests of cophylogenetic hypotheses often employ tree reconciliation methods that infer evolutionary processes such as cospeciation, host shifts, duplications, and losses to account for topological incongruence between host and associate phylogenies ( Page 1994). This approach has the power to model the relative contributions of different evolutionary processes to a given phylogenetic pattern, but biologically realistic scenarios become computationally intractable for large numbers of taxa ( Merkle and Middendorf 2005 Ovadia et al. 2011). Genetic algorithms that incorporate dynamic programming to efficiently locate and evaluate samples from an extremely large universe of event-based solutions hold promise in this regard ( Conow et al. 2010).

Here, we extended the application of a genetic algorithm to event-based tree reconciliation analysis for cophylogenetic problems involving >100 taxon pairs and applied randomization tests involving null models to test the codivergence hypothesis on an unprecedented scale. Nearly 200 pairs of interacting fig and fig wasp species were sequenced at 5 fig loci (providing up to a total of 5.5 kb DNA sequence) and 6 wasp loci (up to a total of 5.6 kb). Two supermatrices were assembled. On an average, wasps had sequences from 77% of 6 genes, figs had sequences from 60% of 5 genes, and overall, we generated 850 new DNA sequences for the purpose of this study. Maximum likelihood (ML) analyses of fig and wasp data sets and Bayesian phylogenetic analyses under relaxed molecular clock assumptions enabled the comparison of distance, event-based, and temporal congruence. Inferences from historical biogeography based on our global sample of fig and pollinator clades provided additional insight on the relative roles of dispersal and vicariance with respect to alternative hypotheses of diversification.


Chaitophorus leucomelas

Adult apterae of Chaitophorus leucomelas are elongate oval in shape and the background colour varies from green to yellow. They usually have two dark stripes along the sides which tend to merge on abdominal tergite V (see first picture below), but sometimes the stripes may be divided segmentally (see second picture below), or may be missing altogether. Whatever the pattern, there are (nearly) always broad pale spinal and marginal areas. The antennae are half as long as the body of the aphid, with the terminal process of antennal segment VI 2.7-3.3 times the base of that segment. The fused apical rostral segment (RIV+V) is 0.8-1.2 times the length of the second hind tarsal segment (HTII). Abdominal tergite I is usually free from tergites II-VI (cf. Chaitophorus tremulae, which has abdominal tergite I fused with tergites II-VI). The hind tibiae are without any pseudosensoria (cf. Chaitophorus populeti, which has a small number of pseudosensoria on the hind tibiae even on the adult vivipara). The short truncate siphunculi are dark at least apically (cf. Chaitophorus populialbae, which has entirely pale siphunculi). The cauda is rounded and very pale . The body length of adult Chaitophorus leucomelas apterae ranges from 1.2 to 2.4 mm.

Chaitophorus leucomelas alatae have dark brown dorsal abdominal cross-bands of varying width (see two pictures below) and separate marginal sclerites visible in the images.

The micrographs below show dorsal views of Chaitophorus lecomelas aptera (pale form) and alate (dark form).

The host plant of Chaitophorus leucomelas in Europe is mainly black poplar (Populus nigra) and related species and hybrids, but in North America a wider range of species is colonised. They feed on young shoots of Populus spp. in spring, and later under leaves, in leaves stuck together by moth larvae, or in leaf galls vacated by other insects. The black poplar leaf aphid is commonly ant-attended. Chaitophorus leucomelas is widely distributed in Europe, North Africa and Asia, and has been introduced to South Africa and North and South America.

Biology & Ecology

Life cycle

The overwintering eggs of Chaitophorus leucomelas hatch in spring and the fundatrices develop on the young leaves.

The development rate of this species on black poplar was studied under laboratory conditions by Hintze-Podufal & Thorns (1978). The development rate almost doubled when the temperature was increased from 15 °C to 25 °C. However, the number of progeny at 25 °C was reduced, and the death rate at the higher temperature was greater especially during the first larval stages. The lower life expectancy and lower fecundity at 25 °C were compensated for by the more rapid development at that temperature.

No studies appear to have been carried out to assess the nutritional benefits (or otherwise) of feeding on tissue galled by other insects. It seems likely that availability of soluble nitrogen will be higher in galled leaves, which would be advantageous to Chaitophorus leucomelas enabling it to continue to feed and reproduce in mid-summer when no further young leaves are available.

Oviparae (see picture below) and males are produced in October, and eggs are laid on poplar twigs.

Colour

Chaitophorus leucomelas has unusually variable dorsal markings. The picture below shows part of a colony of this species including three adult apterae. Their colour ranges from very dark to pale.

The dark aptera on the right has the typical markings of Chaitophorus leucomelas with two dark stripes along the sides which merge on the fifth abdominal tergite. The central, virtually all dark, aptera is unusual - Blackman & Eastop (1994) use the characteristic of 'broad pale spinal and marginal areas' in his key, but this would not work on this specimen. The paler aptera on the left would correctly key out to Chaitophorus leucomelas given it has dark siphunculi. All specimens have a noticeably very pale, almost white, cauda.

Coeur d'acier et al. (2014) notes that Chaitophorus leucomelas has exceptionally high intraspecific divergences. It is a species with a large geographic distribution that presents different numbers of chromosomes according to its origin. This suggests that there may be sibling species within this taxon.

Host selection

Insect response to plant surface features is a critical step in host-finding and acceptance of herbivorous insects. The plant surface is usually covered with epicuticular waxes. which are not only involved in water physiology, but also provide resistance to insects. Alfaro-Tapia et al. (2007) looked at the probing behaviour and performance of the aphid Chaitophorus leucomelas on dewaxed and waxed leaves of two poplar hybrids. Laboratory experiments showed that in naturally waxed leaves of the resistant hybrids, aphids devoted less time to probing and more time to non-probing behaviour when compared with their behaviour on susceptible hybrids. These differences were not present when leaves of these hybrids were experimentally dewaxed. A field experiment demonstrated that aphid reproductive performance was affected by hybrid genotype, but not by epicuticular waxes, although a trend of lower performance on dewaxed leaves in both hybrids was apparent.

Barrios-San Martin et al. (2014) investigated host selection and probing behaviour of the poplar aphid testing the hypothesis that poplar resistance to this aphid is associated with the presence of volatiles and secondary plant compounds. Studies on two poplar hybrids with contrasting susceptibilities suggested the involvement of antifeedant factors in the less favoured hybrid as well as a higher abundance of monoterpenes, sesquiterpenes, alkanes and phenols.

Competition / coexistence

Chaitophorus leucomelas is unusual in that it tends to live inside the galls of other insects, indeed often the galls of other aphids. One such aphid gall often used by Chaitophorus leucomelas is that of Thecabius affinis, the poplar-buttercup gall aphid (see picture below).

The images below show an aptera and an alate inside the gall produced by the poplar-buttercup gall aphid on black poplar.

The yellowish immatures are also mostly those of of Chaitophorus leucomelas. They do not produce wax, but are often thinly covered with wax strands which are picked up from the wax produced by Thecabius affinis. The large alate below is a Thecabius affinis that has developed in the gall.

In the case above, Chaitophorus is likely to benefit from the association both from the protection from predators and parasitoids provided by the gall plus wax, and nutritionally through the 'sink effect'. In the case below, Chaitophorus leucomelas were selectively feeding on a bacterial gall caused by Taphrina populina.

There is no wax and little protective covering - the gall is completely open on the underside of the leaf. Hence the only benefit to Chaitophorus is any increase in soluble nitogen resulting from the bacterial gall.

Ant attendance

Chaitophorus leucomelas is sometimes attended by ants which feed on the copius amounts of honeydew excreted by the aphids (see picture below).

Within the genus Chaitophorus, Shingleton et al. (2005) looked at how a morphological trait - the length of the mouthparts - may have promoted the evolution of ant-aphid mutualisms. Considering thirteen species of Chaitophorus, they suggested there was an evolutionary relationship between feeding position, dimensions of the mouthparts, ability to escape, and the risk of predation suffered by an aphid species. Aphid mouthpart length varied with feeding position with petiole-feeding species having longer mouthparts than leaf-feeding species. Differences in mouthpart length influenced an aphid's ability to escape from predators. There was a significant positive relationship between the mean length of a species' mouthparts, and the mean time for that species to escape. Differences in escape ability influence an aphid's susceptibility to predation. There was a positive relationship between mean mouthpart length for a species and the level of predation it suffered. Given these findings, they then used the data to test whether there was a relationship between mouthpart dimension and ant tending. Amongst the Chaitophorus aphids, tended aphids (obligate and facultative) had longer mouthparts than untended aphids.

The mouthpart length of Chaitophorus leucomelas is about 0.30 mm, midway in Shingleton's mouthpart length distribution of Chaitophorus species, and similar to other leaf feeders either facultatively ant tended (Chaitophorus populialbae) or untended (Chaitophorus salijaponicus niger). Both Shingleton et al. (2005) and Blackman & Eastop (1994) class Chaitophorus leucomelas as an obligate myrmecophile. Our own observations cast doubt on this because they do not seem to be attended when living in galls, whether induced by Taphrina or by Thecabius affinis. We would instead class this species as a facultative myrmecophile.

Natural enemies

When the aphids are not attended, we have observed predatory syrphid larvae preying on the colonies (see picture below).

Tomanovic et al. (2009) describes a new species of braconid, Areopraon chaitophori, associated with Chaitophorus leucomelas on Populus.

Other aphids on same host

Chaitophorus leucomelas has been recorded from 22 Populus species.

Blackman & Eastop list 41 species of aphid as feeding on black, or Lombardy poplar (Populus nigra) worldwide, and provide formal identification keys (Show World list). Of those aphid species, Baker (2015) lists 17 as occurring in Britain (Show British list).

Damage and control

Most research on control of black poplar leaf aphid is focused on the use of plant resistance to minimise pest damage. Chaitophorus leucomelas is one of the most important pests of poplar (Populus spp.) plantations in Iran. Yali et al. (2009) assessed reproduction, and life history of Chaitophorus leucomelas on 11 poplar clones belonging to three species, Populus nigra, Populus deltoides and Populus euramericana. They concluded that susceptibility is inherited through Populus deltoides, whereas resistance seems to be inherited through Populus maximowiczii. Thus, Populus maximowiczii hybrids are recommended for commercial or ornamental planting programs in zones where there is a high risk of aphid infestation. Ripka (1999) lists Chaitophorus leucomelas as one of the pests causing the biggest problems on ornamental trees and shrubs in Hungary.

Identifications & Acknowledgements

Whilst we make every effort to ensure that identifications are correct, we cannot absolutely warranty their accuracy. We have mostly made identifications from high resolution photos of living specimens, along with host plant identity. In the great majority of cases, identifications have been confirmed by microscopic examination of preserved specimens. We have used the keys and species accounts of Blackman & Eastop (1994) and Blackman & Eastop (2006) supplemented with Blackman (1974), Stroyan (1977), Stroyan (1984), Blackman & Eastop (1984), Heie (1980-1995), Dixon & Thieme (2007) and Blackman (2010). We fully acknowledge these authors as the source for the (summarized) taxonomic information we have presented. Any errors in identification or information are ours alone, and we would be very grateful for any corrections. For assistance on the terms used for aphid morphology we suggest the figure provided by Blackman & Eastop (2006).

Useful weblinks

References

Alfaro-Tapia, A. (2007). Effect of epicuticular waxes of poplar hybrids on the aphid Chaitophorus leucomelas (Hemiptera: Aphididae). Journal of Applied Entomology 131(7), 486-492. Full text

Barrios-San Martin, J. et al. (2014). Host selection and probing behavior of the poplar aphid Chaitophorus leucomelas (Sternorrhyncha: Aphididae) on two poplar hybrids with contrasting susceptibility to aphids. Journal of Economic Entomology 107(1), 268-276. Full text

Couer d'acier, A. et al. (2014). DNA barcoding and the associated [email protected] Website for the Identification of European aphids (Insecta: Hemiptera: Aphididae). PLOS ONE 9(6): e97620. doi:10.1371/journal.pone.0097620 Full text

Hintze-Podufal, C. & Thorns, H.J. (1978). Development of poplar aphids (Chaitophorus leucomelas) on leaf discs at 15 deg C and 25 deg C. Journal of Applied Entomology 87(1-4), 388-392. Abstract

Ramirez, C.C. (2004). Differential Susceptibility of Poplar Hybrids to the Aphid Chaitophorus leucomelas (Homoptera: Aphididae). Journal of Economic Entomology 97(6), 1965-1971. Full text

Ripka, G. (1999). Növénykárosító ízeltlábúak a díszfákon és a díszcserjéken: pajzstetvek, levéltetvek, atkák. (Arthropod pests of ornamental trees and shrubs: scale insects, aphids, mites). Növényvédelem 35(12), 623-626. Abstract

Shingleton, A.W. et al. (2005). The origin of a mutualism: a morphological trait promoting the evolution of ant-aphid mutualisms. Evolution 59(4), 921-926. Full text

Tomanovic, et al. (2009). Areopraon chaitophori n. sp. (Hymenoptera: Braconidae:Aphidiinae) associated with Chaitophorus leucomelas Koch on poplars, with a key for European Areopraon Mackauer species. ann. soc. entomol. Fr. 45(2), 187-192. Full text


Insects and Ticks > Black Flies

Black flies, known also as "buffalo gnats" and "turkey gnats," are very small, robust flies that are annoying biting pests of wildlife, livestock, poultry, and humans. Their blood-sucking habits also raise concerns about possible transmission of disease agents. You are encouraged to learn more about the biology of black flies so that you can be better informed about avoiding being bitten and about their public health risk.

Are Black Flies a Public Health Risk?

Black flies can be annoying biting pests, but none are known to transmit disease agents to humans in the U. S. However, they transmit one parasitic nematode worm that infects humans in other regions of the world. Onchocerca volvulus causes a significant human disease known as onchocerciasis or "river blindness" in equatorial Africa and mountainous regions of northern South America and Central America.

The bites of black flies cause different reactions in humans, ranging from a small puncture wound where the original blood meal was taken to a swelling that can be the size of a golf ball. Reactions to black fly bites that collectively are known as "black fly fever" include headache, nausea, fever, and swollen lymph nodes in the neck.

In eastern North America, only about six black fly species are known to feed on humans. Several other species are attracted to humans, but they typically do not bite. However, the non-biting species fly around the head and may crawl into the ears, eyes, nose, or mouth, causing extreme annoyance to anyone engaged in outdoor activities.

Black flies can be found throughout most of the U. S., but their impact on outdoor activities varies depending on the specific region and time of year. For example, in parts of the upper Midwest and the Northeast, black fly biting can be so extreme, especially in late spring into early summer, it may disrupt or prevent outdoor activities such as hiking, fishing, and kayaking.

Besides being a nuisance to humans, black flies can pose a threat to livestock. They are capable of transmitting a number of different disease agents to livestock, including protozoa and nematode worms, none of which cause disease in humans. In addition to being vectors of disease agents, black flies pose other threats to livestock. For example, when numerous enough, black flies have caused suffocation by crawling into the nose and throat of pastured animals. On rare occasions, black flies have been known to cause exsanguination (death due to blood loss) from extreme rates of biting. Saliva injected by biting black flies can cause a condition known as "toxic shock" in livestock and poultry, which may result in death.

How Many Types of Black Flies Are There?

Black flies are true flies (Order Diptera) in the family Simuliidae, which includes more than 1,700 species worldwide. In North America, 255 species in 11 genera have been identified, but additional species remain to be discovered and named. Very little is known about black flies in Indiana, and there are no estimates of the number of species in the state. For perspective, 12 species have been documented in Illinois, while over 30 species have been documented in both Minnesota and Wisconsin, where black fly habitats are more abundant.

How Can I Recognize an Adult Black Fly?

Black flies range in size from 5 to 15 mm, and they are relatively robust, with an arched thoracic region (Figure 1). They have large compound eyes, short antennae, and a pair of large, fan-shaped wings. Most species have a black body, but yellow and even orange species exist.

What Is the Life Cycle of Black Flies?

Black flies undergo a type of development known as "complete metamorphosis" (Figure 2). This means the last larval stage molts into a non-feeding pupal stage that eventually transforms into a winged adult. After taking a blood meal, females develop a single batch of 200-500 eggs. Most species lay their eggs in or on flowing water, but some attach them to wet surfaces such as blades of aquatic grasses.

The length of time it takes an egg to hatch varies greatly from species to species. Eggs of most species hatch in 4-30 days, but those of certain species may not hatch for a period of several months or longer. The number of larval stages ranges from 4-9, with 7 being the usual number. The duration of larval development ranges from 1-6 months, depending in part on water temperature and food supply. The life cycle stage that passes though winter is the last stage larva attached underwater to rocks, driftwood, and concrete surfaces such as dams and sides of man-made channels.

Figure 2. Black fly life cycle. (Illustration by: Scott Charlesworth, Purdue University,
based in in part on Peterson, B.V., IN: IN: Manual of Nearctic Diptera, Volume 1)

The pupal stage is formed the following spring or summer, typically in the same site as the last stage larva, but may occur downstream following larval "drift" with the current. Adults emerge from the pupal stage in 4-7 days and can live for a few weeks. Adults of most species are active from mid-May to July. The number of generations completed in one year varies among species, with some having only one generation, but most species that are major pests complete several generations per year.

Black fly larvae and pupae develop in flowing water, typically non-polluted water with a high level of dissolved oxygen. Suitable aquatic habitats for black fly larval development vary greatly and include large rivers, icy mountain streams, trickling creeks, and waterfalls. Larvae of most species typically are found in only one of these habitats.

Larvae remain attached to stationary objects in flowing water, held on by silken threads extruded from glands located at the end of the bulbous abdomen. Depending on species, mature larvae range from 5-15 mm in length and may be brown, green, gray, or nearly black in color. They possess a large head that bears two prominent structures known as "labral fans" that project forward (see Figure 2). Labral fans are the primary feeding structures, filtering organic matter or small invertebrates out of the water current.

Pupae remain attached to stationary objects in flowing water as well. They typically are orange and appear mummy-like because the developing wings and legs are tightly attached to the body. Pupae of many species produce a delicate, silken "cocoon" of varying density, weave, and size that partially or nearly entirely encloses them other species produce hardly any cocoon at all.

What Should I Know About the Feeding Habits of Adult Black Flies?

It is estimated that females of 90% of the black fly species require a blood meal for the development of eggs. Those of most species feed on mammals, while others feed on birds. Females of some black fly species feed on only one host, whereas others are known to feed on over 30 different host species. No North American species feed exclusively on humans. Male black flies are not attracted to humans, and their mouthparts are not capable of biting.

Females of most species of black flies feed during the day, usually biting on the upper body and head. Unlike certain species of mosquitoes and biting midges, black flies do not enter human structures to seek blood meals.

Do Humans Contribute to Black Fly Problems?

Human activities can lead to an increase in black fly numbers in an area. Structures such as concrete dams and concrete-lined stream channels provide excellent developmental sites for larvae and pupae of certain black fly species. In addition, the restoration of polluted streams, especially in New England, has increased the dissolved oxygen content of streams and created suitable larval habitat for some of our most important pest species.

What Should I Know About Controlling Black Flies?

Control of black flies is difficult, typically aimed at the larval stages, and usually involves aerial applications of insecticides or physically altering the habitat of pest species. The most effective control programs are conducted by state agencies or by professional pest control companies contracted by the state. Any effect is limited in duration, however, in large part because females of pest species are capable of flying long distances from the larval developmental site, and they soon re-infest treated areas.

There is little that an affected homeowner or person engaging in outdoor activities can do to control black flies. For personal protection, it is best to avoid peak periods of black fly activity. Information pertaining to the predicted "black fly season" in a particular area often can be obtained by contacting a local Cooperative Extension office. When venturing outdoors in infested areas, apply an insect repellent containing DEET, wear protective clothing, and minimize openings such as buttonholes through which black flies crawl in an attempt to feed. Outdoor activities in heavily infested areas may require the wearing of fine-mesh head nets, similar to those worn by beekeepers.

Where Can I Find More Information on Black Flies?

A recent (2002) textbook by G. Mullen and L. Durden, Medical and Veterinary Entomology, has an excellent chapter devoted to black flies that covers biology, behavior, medical and veterinary risk, and information on personal protection and approaches to black fly control.


Insect identifier App by Photo, Camera 2020

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Insect identification make up the great biodiversity of the earth. There are several million insect species, and entomologists have divided them into a reasonable number of units called "orders." The members of each insect order come from a common ancestor, have similar structural features, and have certain biological characteristics.

All insect orders are not the same number of species Some orders have only a few hundred species, others more than 100,000. The range of structural features and biological characteristics tends to be wider among the higher-ranking species.

Insect identifier give Predictions on the biology, behavior and ecology of an insect can be made as soon as you know your order. But how do you know which order an insect belongs to? Insects can be identified in several ways. Comparing a specimen to a book of images of identified insects is one possibility. Using a printed key is another way. This Lucid-based key combines the benefits of these methods and adds a new dimension of simplicity and performance to the identification process.

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Pests & Diseases affecting chilli peppers

So, you've bought (or saved) your seeds, carefully planted them and provided the optimum growing conditions. However danger lurks in every corner of the garden with a whole host of beastly pests and diseases ready to indiscriminately strike down your plants at a moments notice.

In general there are two types of factors which can bring death and destruction to your beloved chile plants - living (biotic) and nonliving (abiotic) agents. Living agents incude insects, bacteria, fungi and viruses. Nonliving agents include extremes of temperature, excess moisture, poor light, insufficient nutrients, poor soil pH and air pollutants.

This chilemans guide aims to provide an overview of some of the more common living agents that can infect your Chile plants to help you identify 'the enemy' and provide you with some ammunition to fight the problem. After all thechileman wants your plants to have a long and healthy life and produce a bountiful harvest of lovely chiles.

Know Your Enemy!

Unfortunately there are a whole host of pests & diseases that can infect Chile pepper plants. Thankfully, only a few are common in the UK with most more of a problem in hotter climates such as the Caribbean and Americas. Although most insects are more of an irritation than a terminal problem causing only localised damage, it is the diseases that they can carry which can do the real damage.

A study by Green, S. K. and Kim, J. S (1991), found that more than half of known viruses are transmitted by aphids (greenfly). Thrips, mites, whiteflies, beetles and nematodes transmit others. Some of the more serious problems such as Bacterial Leaf Rot and Tobacco Mosaic Virus (TMV) are transmitted by direct contact with infected plants, soil or garden tools with others transmitted through mechanisms not yet understood.

Diseased plants can exhibit a variety of symptoms, making diagnosis extremely difficult. Common symptoms include abnormal leaf growth, colour distortion, stunted growth, shrivelled plants and damaged pods. Although pests & diseases can cause considerable yield losses or bring death to your plants, none are believed to directly affect human health.

Prevention is better than the cure

As a general rule, most pests and diseases cannot be completey eradicated, but they can be managed and controlled to minimise the 'collateral' damage. Once a problem has taken hold it is often very difficult to control.

To manage potential problems, early identification, correct diagonsis and the swift implementation of preventitive methods should allow you to get on top of most problems before serious damage if inflicted.

However, for the sake of the environment before automatically reaching for the nearest bottle of poison, there are several much friendlier and easy organic strategies which can be deployed, particularly for controling insects. Unfortunately, the more serious viral & fungal infestations may require Chemical Warfare to be deployed. However, always read the instructions on the bottle carefully and take precautions when using chemical agents.

Organic Strategies for Managing Pests

1. Learn to tolerate some damage: Most healthy Chile plants can tolerate some damage without suffering serious long-term problems or yield reduction. Munched leaves/ damage pods can easily be removed to maintain the attractive appearance of your plant.

2. Introduce the 'Good guys': Aphids feeding in the spring can alarm many Chile growers. Introducing natural predators such as Ladybirds, Parasitic wasps and Lacewings will help clean up most local infestations in a month or so.

3. Hand pick/Hunt down: Hunting down snails and slugs and 'disposing of them' can be a highly satisfying exercise particularly if the little blighters have already struck your prized plants. Night time 'slug hunts' during wet weather can be particularly productive.

4. The Water Hose: A strong water hose will temporarily dislodge flies, aphids and other pests from mature plants. However be careful not to saturate or damage your plants and avoid this using method on young seedlings.

5. Remove diseased plants or plant parts: Simply removing and disposing of badly damaged plants can help reduce the problem and prevent is spreading to adjacent plants.

6. Crop Rotation: This is particularly important strategy for tackling soil borne pathogens such as Verticillium Wilt and root rot.

7. Grow pest resistant & pest tolerant plants: Many hybridised varieties, particularly some of the newer sweet pepper varieties have been developed to give specific resistance to diseases such as Tobacco Mosaic Virus (TMV) and Bacterial Leaf Spot.

8. Innoculate: When growing in pots it is likely that sterilized soil has been used. Sterile soil is ripe for colinisation by many forms of bacteria, fungi, & insects. It is quite likely that the first colinization will not be beneficial. However, just as you can buy yogurts containing beneficial bacteria from the supermarket, you can also buy beneficial bacteria for your soil (though it is a little bit more difficult to get hold of). Beneficial Mycorhizzal fungi is also available, and is starting to become popular in many on-line shops. It may also be useful, depending upon the scope of your growing conditions, to introduce beneficial soil dwelling predatory insects. Introducing your own symbiotic bacteria, fungi, and insects limits the likelihood of colonisation by parasitic forms. In addition to aiding growth of the chile plant, & providing tolerance to environmental stresses, many forms of bacteria and Mycorhizzal fungi are also thought to innoculate the chile plant from diseases and viruses. In addition, they are helpful at reducing the conditions that make these diseases and viruses possible.

What's the Problem? - A Quick Reference Guide

Unless you are an expert taxonomist or have easy access to a laboratory, the correct diagnosis of the problem is probably the most difficult (and critical) factor in your battle with the enemy as a whole host of problems can display similar very symptoms. The following guide will hopeful help you narrow down the problem.

The Leaves:

Yellowing
- see the sections on Aphids, Whitefly, Nematodes and Verticillium Wilt
- could also be caused by a Nitrogen or Magnesium deficiency, mineral deficiency, or excessive watering

Browning
- see Bacterial Leaf Spot and Phytopthora blight
- could also be caused by excessive nitrogen.

Curling/distortion
- see Aphids, Thrips, Spider mites and Viruses

Holed
- see slugs & snails and flea beetles

Scorched
- see sunscald
- could be caused by Chemical or fertiliser burns

Spots/Blotches
- see Bacterial Leaf Spot, Cercospora Leaf Spot Powdery Mildew, Phytopthora blight and viruses
- could also be caused by chemical injury

The Plants:

Browning Stems
- see Bacterial Leaf Spot and Phytopthora blight
- could also be caused by insufficient watering

Wilting
- see Verticilllium wilt, Bacterial Wilt & Phytopthora blight
- could also be caused by too little/too much watering

Plants Falling Over
- could be caused by waterlogged soil, insufficient plant support or poorly develop roots

Slow growth
- likely to be caused by inadequate light, poor soil, low temperatures. Note some Chile species particularly the Chinese are notoriously slow growing

The Pods:

Holes
- see slugs & snails and pepper maggots
- Birds and animals are also partial to the occasional chile pod (animals tend to avoid all but the mildest chile pods - though they might take a test nibble).

Spots/discolouration
- see Anthracnose, Bacterial leaf spot, Blossom End Rot, Phytopthora blight, Grey Mold and thrips
- could also be caused by sunscald or nutrient deficiencies

Distortion
- see Thrips, Spider mites and viruses. Poor Pollination can also cause this problem

Soft Rot
- see Bacterial Soft Rot and Grey mold

Failure to Ripen
- insufficient ripening time likely to be the problem

Insect Pests

The insects most likely to 'enjoy' your chile plants are slugs & snails, aphids (greenfly/blackfly), pepper maggots,whitefly and nematodes. Flea beetles, cutworms, hornworms, thrips, spider mites and leafminers are less common. To control insect problems, regular inspection is again the key to success.

Slugs & Snails are probably the number 1 enemy of gardeners, these little devils can quite happily turn one of your prize specimens into a swiss cheese practically over night before sliding back to there hideaways, leaving you to wonder what happened. Thankfully, most slugs and snails leave behind one piece of incriminating evidence which helps to both diagnose the problem and track them down, a trail of slime! Slugs are hermaphrodites (they can mate with themselves) and can produce dozens of eggs several times a year. The egg clusters look like little piles of whitish jelly and hatch anywhere from 10 days 28 days. 'Dispose' of any slugs and eggs wherever you find them.

Regular Slug hunts are the best course of action. Container gardening, the use of Copper tape/matting (placed around the plant) and even garlic oil has been used by gardeners with some success.

Aphids (Greenfly/BlackFly) are one of the commonest and most annoying all garden insects. They are particularly attracted to young tender shoots, sucking your plants dry of sap causing shoots and leaves to become distorted. Plants grown indoors and away from natural garden predators can be particularly prone to infestations. Image © Virtualpepper

Small infestations are relatively easy to control. One method is to introduce natural predators to do the job for you. A second is to attract them away from your like darlings by planting Marigolds (tagetes and calendula) close by. Marigolds are a feeding favourite of the aphids and the theory goes that they will be much more interested in the Marigolds than your Chile plants.

Other friendly ways of controlling aphids include rubbing them off with your fingers or spraying them with a very diluted soap solution, about one teaspoon of fairy Liquid pure soap (as near to 100% fatty acids as you can get - avoid antibacterial, perfumed, & detergent based soaps) to a couple of litres of water. More severe infestations are more troublesome and it may be better to isolate the plants to prevent the problem spreading to your other plants. Unfortunately, spraying severely infested plants will provide only temporary relief and may simply just shift the aphids from one plant to another.

Flea beetles are about 2mm long, shiny in appearance with enlarged hind legs which enable them to jump. Adult flea beetles feed on the undersides of young leaves leaving small pits or irregularly shaped holes. Larvae live primarily in the soil and feed on roots, but cause little damage.

Ensure rapid germination and development of seedlings so that they grow through this vulnerable stage quickly. Flea beetles feed at the height of the day, and they don't like to get wet. Giving them a lunchtime shower can reduce the problem.

Pepper Maggots are whitish yellow, pointed at the head end and 0.5in longwhen fully grown. The maggots feed on the core inside of the pods which causes damaged peppers to turn red prematurely and rot.

Check pods for small puncture holes and destroy and infected pods. Rotting pods will attract other flies if left on the plant.

Root Knot Nematodes are microscopic, eel-like roundworms that live in the soil and feed on roots. Root damage reduces the plants ability to take up water and vital nutrients. Symptoms vary with plant age and the severity of the infestation, but include wilting, nonproductive plants and development of characteristic knots on the plant's roots which can vary in size from smaller than a pinhead to larger than a pea. The problem can be particularly severe in sandy soils.

Crop rotation and adding organic matter to sandy soils can help reduce the impact of nematodes. The best method of control is to plant resistant varieties (often indicated by an N on the seed packet) like California Wonder & Charleston Belle.

Spider Mites can be a serious problem particularly during periods of hot, dry weather. They feed on the underside of leaves and to the naked eye, look like moving dots. When infestation is high, the leaves will have webs on them if uncontrolled, these mites can kill a plant. Infected leaves often curl downwards and leaves are speckled in appearance, as though covered with hundreds or thousands of pale yellow dots. A simple technique for identifying mites is to tap an infected leaf over a piece of white paper. Wait a few seconds and watch for movement.

Red spider mites breed in hot and dry places. If you can increase the humidity around the plant you decrease the pest's reproduction rate. Dampen down infected areas. For house plants a short holiday somewhere cooler and more humid (the bathroom?) may help get rid of the infestation.

Thrips are numerous in species and all are extremely small. They are very slender and may be white, yellow, brown or black. Affected leaves are often distorted and curl upward. The lower surface of the leaves can develop a silvery sheen that later turns bronze. Damage on pods appears as brown or silver areas near the calyx.

Thrips do not usually need to be controlled as predatory mites insects will normally do the job for you.

Whiteflies are tiny insects (1.5mm long) with broad wings that fly from the plant when disturbed. They suck plant juices from the leaves, causing them to shrivel, turn yellow and drop. Whiteflies also secrete honeydew which can cause foliage to become sticky and coated with a black sooty mold.

Whitefly control is difficult, since only the last (flying) stage of the whitefly lifecycle is vunerable to spraying. Whitefly control is difficult as they have very fast lifecycles. To eliminate this pest frequent spraying is necessary - at least once a week, and for many weeks/months. Good cultural practices, such as removing infected plants, pruning the top new growth, and/or using a mild diluted (fatty acid based) soap solution are possible controls. Perseverance is neededare the best controls.

Bacterial & Fungal Diseases

Anthracnose is caused by the fungi Colletotrichum piperatum and C.capsici and is promoted by warm temperatures, high moisture and poor circulation among the plants. Both sweet and hot peppers varieties are susceptible to this disorder. Although the disease does not seriously affect vegetative growth, it can seriously damage pods. Symptoms appear on both ripe and unripe pods and are characterised by sunken, circular spots that can grow up to 1in in diameter. In moist conditions, pink or yellow spore masses may appear.

Crop rotation and the use of disease-free seed. If the disorder is severe, a fungicide may be needed.

Bacterial Leaf Spot is caused by theseed borne bacterium Xanthomonas campestris pv vesicatoria which also causes bacterial spot in tomatoes and is one of the most serious bacterial disease affecting chiles. The principle sources are infected seed and transplants. Moist conditions encourage disease development.

This disease first appears as small water soaked areas that enlarge upto a quarter inch in diameter. The disease spots have black centres and yellow halos. The spots are depressed on the upper leaf surface, whereas on the lower surface the spots are raised and scab like. Severely spotted leaves will eventually turn yellow and drop off, leaving pods susceptable to sunscald.

Crop rotation and the use of disease-free seed. The use of copper-based fungicides can have some success although excessive use may retard growth and damage plants

Bacterial Soft Rot is caused by bacterium Erwinia carotovora pv carotovora and affects Chile pods. The internal tissue softens before eventually turning into a watery mass with a foul smell. This problem is worst in wet weather because the bacteria are splashed from the ground and onto the fruit. It can also be started by insect injury.

Keep plants off the ground (on greenhouse staging) and controlling insects can help reduce the threat of this disorder.

Bacterial Wilt is caused by the bacterium Pseudomonas solanacearum. The first symptoms start with the wilting of the leaves. After a few days, a permanent wilt of the entire plant results, with no leaf yellowing. You can test for this bacteria by cutting the roots and lower stems look for milky streams of bacteria when they are suspended in water.

The best control is to plant clean seed and transplants and to remove diseased plants.

Cercospora Leaf Spot (Frog Eye) is caused by the fungi Cercospora capsici and is worst under extended warm, wet conditions. This disease in characterised by small brown circular leaf lesions that have a watery appearance. Excessive leaf drop may occur in common infestations.

Clean seed and crop rotation are the best preventative measures against this disease. Good airflow around plants in sheltered areas (greenhouse's) will also help minimise this problem. Fungicides are probably the best solution if the problem is extensive.

Damping-off is caused by poor seed quality, improper planting depth, high salt concentrations, a wet seed beds or severe nutrient deficiencies. Several fungi such as Pythium, Rhizoctonia and Fusarium are associated with this problem. Seedlings fail to emerge (pre-emergence damping-off), small seedlings collapse (post-emergence damping-off), or seedlings are stunted (root rot and collar rot).

To control this problem plant only high-quality seed or vigorous transplants and avoid soil that is poorly drained. Good ventilation reduces surface moisture, and therefore the likelihood of damping off. The use of a fungicide, such as a copper based fungicide, or even just watering with chamomile tea (provides a mild fungicide at normal strength), can reduce the likelihood of damping off further.

Grey Mold is a relatively common problem and is caused by the fungus Botrytis cinerea. Symptoms include a sudden collapse of succulent tissues, such as young leaves, stems, and flowers. Grey powdery spore masses occur on the surface of dead plant tissues.

High humidity favours the disease. Ensuring your plants have good air circulation will helps reduce this problem. A fungicide is probably the best bet if the mold is severe.

Phytophthora Blight (Chile Wilt) is caused by a water borne fugus Phytophthora capsici and is generally observed in wet waterlogged areas. The fungus can invades all plant parts causing at least three separate syndromes: leaf blight, fruit rot, and root rot. It is promoted by warm, wet weather. Plants suffering from this condition often wilt and die, leaving brown stalks and leaves and small, poor-quality fruits. If the fungus enters the roots, the game is unfortunately over as the plants cannot obtain enough water (due to root rot), suddenly wilt, and eventually die. Symptoms of the less serve leaf blight include brown or black spots that may kill a localised portion of the plant. Affected areas are often bordered with a white mold.

Avoid excess watering and poorly drained soil. Fungicides can be used to treat leaf blight and fruit rot. Root rot is usually terminal.

Powdery Mildew is cause by the fungus Leveillula taurica and primarily affects leaves on pepper plants during warm wet conditions. Although the disease commonly occurs on older leaves just before or at fruit set, it can develop at any stage of crop development. Symptoms include patchy, white, powdery growth that can enlarge to cover the entire lower leaf surface Diseased leaves eventually drop off, leaving pods susceptable to sunscald.

Powdery mildew is managed primarily with fungicides. However sprays of sulphur and potassium bicarbonate have been known to have some success.

Verticillium Wilt is caused by soil borne fungus Verticillium dahliae is a soil borne fungi which can infect pepper plants at any growth stage. Cool air and soil temperatures favour it. This problem is particularly hard to pin down, as the symptoms are highly variable. Plants may show a yellowing of leaves and stunted growth. As the disease progresses, the plants can shed leaves and may finally die. If the stem is cut, a brown discoloration may be visible.

Crop rotation is the best control. Neither resistant cultivars nor chemical controls are known.

White Mold is caused by the fungus Sclerotinia sclerotiorum. It causes blighting or rotting of any above or below ground plant parts. At first, the affected area of the plant has a dark green, greasy, or water-soaked appearance. On stems, the lesion may be brown to grey in colour. If the humidity is high, a white, fluffy mold growth may appear.

Controls includes well-drained soil, proper plant spacing, crop rotation, and careful removal of all infected plants as soon as possible. Do not compost or use diseased plants for mulch.

Viral Diseases

Pepper Mosaic & Pepper Mottle Virus (PeMV) is caused when infected aphids and other insects come into direct contact with the plant. Stunted plants, distorted fruit, and yield reduction are all symptoms.

Aphids control and good sanitation practises. Planting resistant varieties is the best way to avoid this problem. Early detection and removal of infected plants helps, but complete control is often difficult.

Tobacco Etch Virus (TEV) is caused when infected aphids and other insects come into direct contact with the plant. Dark green vein bands, leaf distortion and stunted growth are all symptoms. Tabasco Chile plants are particularly susceptible to this disease and often wilt and die.

Aphids control and good sanitation practises. Planting resistant varieties is the best way to avoid this problem. Early detection and removal of infected plants helps, but complete control is often difficult.

Tobacco Mosaic Virus (TMV) is a highly infectious and persistent disease is carried by tobacco in cigarettes and is spread mechanically, by infected hands touching tools or plants. Symptoms can include curling leaves, spotted or mottled fruit, stunted plants and excessive leaf drop.

Smokers should disinfect hands (milk kills TMV) thoroughly before gardening. Growing resistant varieties is the best prevention. Early detection and removal of infected plants helps, but complete control is often difficult. Further Information Sources: The University of Maryland Cooperative extension www.hgic.umd.edu

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