Microbial Insecticides Ziad W Jaradat

Microbial Insecticides  

Insects are the most abundant group of organisms on Earth, and they negatively affect humans in a variety of ways: They cause massive crop damage, They act as vectors of both human and animal diseases.

History of insecticides During the 1940s, a number of chemical insecticides were developed as a means of controlling the proliferation of noxious insect populations.

One of these was the chlorinated hydrocarbon DDT (di chlorodiphenyltrichloroethane). DDT proved to be exceptionally effective in killing and controlling many species of pests by attacking the nervous system and muscle tissue of insects. Other chlorinated hydrocarbons such as dieldrin, aldrin, chlordane, lindane, and toxophene have since been synthesized and applied on a massive scale.

Another class of chemical insecticides is called organophosphates and includes malathion, parathion, and diazinon.

The first generation of organophosphates were developed as chemical warfare agents. Now they are used to control insect populations by inhibiting the enzyme acetylcholinesterase, which hydrolyzes the nerve transmitter acetylcholine. These insecticides disrupt the functioning of motor and brain neurons of the insect.

By early 1960s, over 100 million acres of U.S. agricultural land were being treated annually with chemical insecticides. However, at a latter time, researchers realized that chlorinated hydrocarbon insecticide and organophosphate insecticides had dramatic effects on animals, ecosystems, and humans.

DDT, was found to persist in the environment for 15 to 20 years and accumulates in increasing concentrations through food chains. This bioaccumulation in fatty tissues was having a significant biological impact on many organisms. For example, in North America, many species of birds including peregrine falcons, sparrow hawks, bald eagles, brown pelicans, and double-crested cormorants were severely depopulated.

Drawbacks of chemical insecticides;Targeted insect pest populations be increasingly resistant to treatment with many chemical insecticides

Chemical insecticides were found to lack specificity; consequently, beneficial insects were being killed along with those that were considered to be pests. Some times the natural enemies of the insect pest species were killed more efficiently than the target organisms.Given all the drawbacks associated with the use of chemical inseciticides, alternative means of controlling harmful insects have been sought over the past 20 years.

Using insecticides that are produced naturally by either microorganisms or plants was an obvious choice, Why? Highly specific for a target insect species Biodegradable, Slow to select for resistance.

Researchers can manipulate genes that encode insect pathogenic agents and introduce them in target microbes that can infect these insects.

Insecticidal Toxin of Bacillus thuringiensisA microbial insecticide can be an organism that either produces a toxic substance that kills an insect species or has the capability of fatally infecting a specific target insect. The most studied, most effective, most often utilized microbial insecticides are the toxins synthesized by Bacillus thuringiensis. This bacterium comprises a number of different strains each of which produces a different toxin that can kill certain specific insects, for example;

B. thuringiensis kurstaki is toxic to; lepidopteran larvae including moths, butterflies, skippers, cabbage worm, and spruce budworm.

B. thuringiensis israelensis kills diptera such as mosquitoes and black flies. B. thuringiensis tenebrionis is effective against coleoptera (beetles) such as the potato beetle and the boll weevil. Other B. thuringiensis strains with different toxins that are specific toward certain insects.

 

Mode of actionTo kill an insect pest, B. thuringiensis must be ingested as the contact of the bacterium or the toxin with the surface of an insect has no effect on the target organism. B. thuringiensis is generally applied by spraying, so it is usually formulated with insect attractants to increase the probability that the target insect will ingest the toxin.

Advantage; Because for the toxin to be effective it has to be ingested, this limits the susceptibility of none target insects and other animals to this insecticide.  

Drawbacks; Insects that attack plant roots are less likely to ingest a B. thuringiensis toxin that has been sprayed on the surface of a host plant.

B. thuringiensis toxin can only kill a susceptible insect during a specific developmental stage. It costs 1.5-3 times as much as chemical insecticidesResistance of insects to the toxins produced by these bacteria might occur.

Example of using Bacterial insecticides;

Bacillus thuringiensis sub specs. kurstaki was used as major means of controlling spruce budworm in Canada.Its use was increased from 1% in 1979 to around 74% in 1986 for treating spruce budworm in Canada In other countries, B. thuringiensis kurstaki has been used against tent caterpillars, gypsy moths, cabbage worms, cabbage loopers, and tobacco hornworms.

How Does it Work

The insecticidal activity of B. thuringiensis kurstaki and other strains is contained within a very large structure called the parasporal crystal, which is synthesized during bacterial sporulation. The crystal is an aggregate of one kind of protein that can be dissociated by mild alkali treatment to yield two subunits of 130 kDa each (Fig. 12.1). The parasporal crystal is not the active form of the insecticide; rather, it is a protoxin, a precursor of the active toxin.

When the para sporal crystal is ingested by a target insect, the protoxin is activated within its gut by the combination of alkaline pH (7.5 to 8.0) and specific digestive proteases, which converts the protoxin into an active form with 68 kDa (Fig. 12.1). When the toxin changes to its active form, it inserts itself into the membrane of the gut epithelial cells of the insect and creates an ion channel through which excessive loss of cellular ATP occurs (Fig. 12.2). About 15 min after the channels forms cellular metabolism ceases, the insect stops feeding, becomes dehydrated, and eventually dies.

Two things make this process very specific;

1. The need for an alkaline media2. The need for specific proteases

B. thurigien is kurstaki is applied by spraying approximately 1 .3 to 2.6 X 108 spores per square foot of the target area at the peak of the larval population of the target organism.The crystal is short lived as it breaks down after exposure to sunlight so it is appropriate to spray it in cloudy days.

Toxin Gene IsolationIsolate and characterize the protoxin gene(s).determine whether the toxin genes are located on a plasmid or on the chromosomal DNA

Test for plasmid-borne toxin genes; a toxin producing strain can be conjugated to a strain that lack the insecticidal activity. If the latter strain acquires the ability to synthesize the insecticidal toxin, then the toxin gene(s) is most likely present on a plasmid because the transfer of chromosomal DNA during conjugation is a rare event.

Isolation of the protoxin encoding DNA sequence

Total cellular DNA is isolated and separated into plasmid and chromosomal DNA using cesium chloride gradient centrifugation.A clone bank is constructed from the chromosomal DNAWhen toxin gene is plasmid encoded, the plasmid can be further fractionated by sucrose gradient to separate the plasmids to different sizes. Figure 12.3Bacillus thuringiensis kurstaki contains an insecticidal protoxin gene on one of 7 different plasmids (2, 7.4, 8.2, 14.4, 45 and 71 kb in length.

To determine which B. thuringiensis kurstaki plasmid carries the protoxin gene, following sucrose gradient centrifugation, the plasmid DNA sample is divided into three fractions

Small (2.0 kb)Medium (7.4, 7.8, 8.2, and 14.4 kb)Large (45 and 71 kb)

The fraction with the small plasmid (2.0 kb) is discarded, because this plas mid is too small to encode a protein equivalent to the 130-kDa protoxin.

The medium and large plasmid fractions are each partially digested with the restric tion enzyme Sau3AI and then ligated into the BamHI site of plasmid pBR322.These clone banks were trans formed into E. coli and then screened immunologically using the following procedure:

• Colonies are transferred from agar plates to a nitrocellulose membrane.

• The transferred colonies are partially lysed with organic sol vents.

• All available sites on the membrane to which primary and sec ondary antibodies could bind are blocked by treating the membrane with bovine serum albumin.

• The bovine serum albumin-treated membranes are treated with rabbit antiserum that contains antibodies against the insecticidal toxin.

• The membranes are washed to remove unbound antibodies and then treated with 251-labeled S. aureus protein A, which binds to the Fc portion of the bound antibodies.

• Spots on the membrane corresponding to colonies that actively synthesize the insecticidal toxin are visualized by autoradiography.

Using the protoxin gene, the 71-kb plasmid of B. thuringiensis. kurstaki was found by DNA hybridization to encode the toxin gene.

Genetic Engineering of B. thuringiensis Strains

First step is to isolate and sequence the toxin gene, then to obtain the amino acid sequence of toxin. When the amino acid sequence was compared for other toxins they all showed a common toxin domain.

It was found that the whole gene is not necessary for the toxin to have its insecticidal activity, rather a portion, a chemically synthesized coding sequence or of course the whole gene can be used for further genetic manipulation

The B. thuringiensis protoxin protein is only synthesized during the sporulation phase of growth therefore,

It might therefore be advantageous to have the toxin gene transcribed and translated during vegetative growth.

This would permit the toxin to be synthesized by a continues fermentation process which decreases the cost of toxin production 

However, when a continuously active (constitutive) promoter form tetracycline resistant gene was introduced into B. thuringiensis the active toxin protein was produced continuously through out the whole growth cycle ( spore and vegetative phases) Figure 12.4, and even the toxin synthesis occurred in B. thuringiensis defective in sporulation gene.

Therefore, under these conditions the toxin will be produced in high quantities.Many crops might be attacked by different insect species, therefore, it would be advantageous if we could create microbial insecticides with broad spectrum of target insects. This could be done by;Transferring a gene against one insect into a B. thuringiensis that already produced another toxin against another insect

Fusing portions of two different species-specific toxin genes to one another so that a unique hybrid toxin is produced and is effective against these to different insects.

Testing whether the insect target can be widened

Toxin genes were taken from B. thuringiensis subspecies aizawai and tenebrionis and used to make one construct (vector)The vector was introduced onto B. thuringiensis subspecies aizawai, tenebrioinis, kurstaki, israelensisAll the transformed strains were tested for their toxicity toward three different larvae.Table 1 shows the results of this experiment.

Some times B. thuringiensis might not be the best bacteria to be sprayed to combat an insect, so the toxin genes must then be introduced into anther suitable vector such as Caulobacter crescentus or cyanobacteria.

Another example is that some insects attack the roots of the plants which makes them inaccessible to the B. thuringiensis –based insecticidal therefore, it is possible to introduce the toxin gene from the Bacillus to a bacteria that colonizes close to the roots. The engineered bacteria could be introduced into the soil so they;

release the toxin close to the roots Since they colonize the soil, they keep producing the toxin continuously thus obviating the need for continuous spraying.

Example; the B. thuringiensis kurstaki insecticidal toxin was introduced into the chromosomal DNA of Pseudomonas fluorescens that colonize corn roots.

 

1. A transposon Tn5 element that had been cloned into a plasmid was genetically modified by altering portions of its left and right borders and deleting its transposase. Such an altered Tn5 element cannot be excised from the plasmid, even by exogenous transposase. Transposes is an enzyme that is encoded by transposon gene and facilitates the insertion or excision of the transposon form a chromosomal site.  2. An isolated B. thuringiensis subsp. kurstaki insecticidal toxin gene was spliced into the middle of the altered Tn5 element on the plasmid and placed under the control of a constitutive promoter.

3. A wild-type Tn5 element was transposed into the chromosome of the root-colonizing strain of P. fluorescens.4. The plasmid carrying the altered Tn5 element with the insert toxin gene was introduced into the host bacterium that had the integrated wild-type Tn5 element. 5. Homologous recombination by means of a double crossover between the nontransposable Tn5 element on the plasmid that carries the toxin gene and the chromosomally integrated wild-type Tn5 led the integration of the altered Tn5 with the toxin gene into the chromosomal DNA, with the concomitant loss of the wild-type Tn5 element.

This form of engineering controls two things; toxin gene is unlikely to be lost either during the large-scale laboratory growth or after release of the engineered microorganism to the environment.

Probability of transfer of the toxin gene to other microorganisms in the environment is very low.

Baculoviruses as Biocontrol Agents Baculoviruses are rod-shaped, double-stranded DNA viruses can infect and kill a large number of different invertebrate organisms. Sub groups of this viral family are pathogenic to several classes of insects including Lepidoptera, Hymenoptera, Diptera, Neuroptera, Trichoptera Coleoptera, and Homoptera. Therefore, some of the baculoviruses are important for the control of certain pests and thus are registered as pesticides. Problems!!!These viruses kill insects slowly within days or weeks.

Solution !!!Enhance the virulence of the virus by introducing foreign genes that will severely impair or kill the target insect such using the gene that disrupts the cell cycle of the insect. During the insect development, larva juvenile hormone is needed for the metamorphosis to happen, now this hormone is degraded by the action of the juvenile hormone estrase which is an enzyme that inactivate the juvenile hormone. Therefore, an increase in the production of this hormone will interrupt the insect life cycle and leads to its death.

 The gene for the juvenile estrase was purified from the insect Heliothis virescens ( tobacco budworm) and the coding sequence was isolated from the cDNA library and inserted into the genome of a baculovirus under the control of the baculovirus transcription signals. When this virus was fed to target insects, the larva feeding and growth was severely limited relative to the control.

Problems of this method; This method is specific for the control of the insects at this larval stage only so it has a limited effect. Therefore, the incorporation of a toxin gene in the genome of the virus that will be transcribed and translated during the viral normal cycle in the insect would help in killing the insect any time regardless of its developing stage.

Example, the gene that encodes the insect specific neurotoxin produced by the North American scorpion was cloned into a baculovirus and was tested against target insects and found to decrease the damage to crops by 50% . The cost of propagating this virus is high