Category: Uncategorized

 

Insect Orders

Locate each insect order in the word search puzzle and then make a list of the common names for the insects in each order.

 

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Mark S. Fradin, M.D., and John F. Day, Ph.D.Insect-transmitted disease remains a major source of illness and death worldwide. Mosquitoes alone transmit disease to more than 700 million persons annually.1 Malaria kills 3 million persons each year, including 1 child every 30 seconds. Although insect-borne diseases currently represent a greater health problem in tropical and subtropical climates, no part of the world is immune to their risks. In the United States, arboviruses transmitted by mosquitoes continue to cause sporadic outbreaks of eastern equine encephalitis, western equine encephalitis, St. Louis encephalitis, and La Crosse encephalitis. In the fall of 1999, West Nile virus, transmitted by mosquitoes, was detected for the first time in the Western Hemisphere. In the New York City area, 62 persons infected with West Nile virus were hospitalized, and 7 persons died.The Centers for Disease Control and Prevention estimates that more than 2000 persons were infected with West Nile virus in the year 2000. The virus has now been detected in 27 states, and it is anticipated that it will continue to spread unabated across the United States during the next few years.

Protection from arthropod bites is best achieved by avoiding infested habitats, wearing protective clothing, and using insect repellent. In many circumstances, applying repellent to the skin may be the only feasible way to protect against insect bites. Given that a single bite from an infected arthropod can result in transmission of disease, it is important to know which repellent products can be relied on to provide predictable and prolonged protection from insect bites. Commercially available insect repellents can be divided into two categories — synthetic chemicals and plant-derived essential oils. The best-known chemical insect repellent is N,N-diethyl-m-toluamide, now called N,N-diethyl-3-methylbenzamide (DEET). Many consumers, reluctant to apply DEET to their skin, deliberately seek out other repellent products. We compared the efficacy of readily available alternatives to DEET-based repellents in a controlled laboratory environment.

Methods

Product Selection

In January 2001, we purchased a total of 16 products for testing, choosing repellents with national, rather than local, distribution (Table 1). Seven widely available botanical repellents were included in the study. Multiple concentrations and formulations of DEET are readily available. We chose and tested three DEET-based repellents (ranging from 4.75 to 23.8 percent DEET) that we believe represented the range of commonly purchased repellents in the United States. We also tested a controlled-release 20 percent DEET formulation to determine whether it had a longer duration of action. The only synthetic repellent containing IR3535 (ethyl butylacetylaminopropionate) that is available in the United States and three wristbands impregnated with either DEET or citronella were also tested. Finally, we tested the efficacy of a proprietary moisturizer that is commonly believed to have repellent effects.

Testing Methods

The duration of protection provided by each product was tested by means of arm-in-cage studies, in which volunteers insert their repellent-treated arms into a cage with a fixed number of unfed mosquitoes, and the elapsed time to the first bite is recorded. Testing of repellents is usually conducted either in a laboratory or at outdoor field sites. Conducting such studies indoors makes it possible to reduce potential confounding variables, such as wind speed, temperature, humidity, density of the mosquito population, the level of the mosquitoes’ hunger, and the species of the mosquitoes, that can make it difficult to interpret comparisons among products made in outdoor-field trials. We conducted our tests with a low density of mosquitoes per cage rather than a high density (some studies use more than 250 mosquitoes per cage) because the low-density environment more accurately reflects the typical biting pressures that are encountered during most outdoor activities.

For each test, 10 disease-free, laboratory-reared Aedes aegypti female mosquitoes that were between 7 and 24 days old were placed into separate laboratory cages measuring 30 cm by 22 cm by 22 cm. A batch of 10 mosquitoes that had not been exposed to the repellent being tested was used for each arm insertion. Mosquitoes were provided with a constant supply of 5 percent sucrose solution. Cages were placed in a walk-in incubator measuring 2.2 m by 2.2 m by 2.2 m, in which the temperature was maintained at 24 to 32°C, the relative humidity at 60 to 70 percent, and the light–dark cycle at 12 hours of light followed by 12 hours of darkness.

Fifteen volunteers (5 men and 10 women) were recruited from the staff of the Medical Entomology Laboratory at the University of Florida. The study was reviewed and approved by the institutional review board of the University of Florida, and subjects gave written informed consent before participating.

As repellents were purchased, they were labeled sequentially from 1 to 16. A random-number generator was then used to determine the order in which the products would be tested on each subject. A total of 720 individual tests were conducted, with each repellent being tested three times on each subject. Most subjects only completed one test per day. The average time to completion of all three tests was 10.2 days. In the case of repellents that were identified as very short-acting in the initial test, subjects were permitted to conduct all three tests of the repellent in a single day, washing the skin with an unscented soap before each application of the repellent. Subjects did not test more than one repellent product on a single day. No information on the likely duration of action of each repellent was provided to subjects before they began their tests.

Before each test, the readiness of the mosquitoes to bite was confirmed by having subjects insert their untreated forearm into the test cage. Once subjects observed five mosquito landings on the untreated arm, they removed their arm from the cage and applied the repellent being tested from the elbow to the fingertips, following the instructions on the product’s label. After the application of the repellent, subjects were instructed not to rub, touch, or wet the treated arm. Repellent-impregnated wristbands were worn on the wrist of the arm being inserted into the cage. Subjects were provided with a standardized log sheet to ensure accurate documentation of the duration of exposure and the time of the first bite. The elapsed time to the first bite was then calculated and recorded as the “complete-protection time” for that subject in that particular test.

Subjects were asked to follow the testing protocol shown in Figure 1. Subjects conducted their first test of each repellent by inserting the treated arm into a test cage for one full minute every five minutes. If they were not bitten within 20 minutes, then the arm was reinserted for 1 full minute every 15 minutes, until the first bite occurred. On the basis of this initial complete-protection time, the subject’s next two tests of that particular repellent were conducted as follows: if the repellent had initially worked for less than 20 minutes, the subject placed his or her arm in the cage for 1 minute every 5 minutes; if the repellent had initially worked for 20 minutes to 4 hours, the subject placed his or her arm in the cage for 1 minute every 15 minutes; and if the repellent had initially worked for more than 4 hours, the subject placed his or her arm in the cage for 1 minute every hour (up to 4 hours). If a repellent was still working after 4 hours, then the subject continued to place his or her arm in the cage every 15 minutes thereafter, until the first bite occurred. If at any point during testing, subjects noted mosquitoes landing but not biting (a behavior that typically occurs when the efficacy of a repellent begins to wane), then the intervals between insertions were decreased to five minutes.

FIGURE 1

Discretionary funds from the State of Florida were used to support this study. We received no financial support from industry, including the manufacturers whose products were tested in the study. Data analysis was performed within the Florida Medical Entomology Laboratory at the University of Florida, without input from any outside sources.

Statistical Analysis

Two-way analysis of variance (involving two factors, subject and repellent) followed by Tukey’s tests was used to compare the mean complete-protection time for the 16 tested repellents. All P values are two-sided; a P value of less than 0.05 was considered to indicate statistical significance.

Results

Of the products tested, those containing DEET provided the longest-lasting protection (Table 1). The complete-protection times of DEET-based repellents correlated positively with the concentration of DEET in the repellent. The formulation containing 4.75 percent DEET provided an average of 88.4 minutes of complete protection; the formulation containing 23.8 percent DEET protected for an average of 301.5 minutes. There was a statistically significant difference in complete-protection time between each DEET-based repellent and the product with the next higher concentration of DEET (P<0.001 for all comparisons). The controlled-release formulation we tested did not prolong the duration of action of DEET. The alcohol-based product containing 23.8 percent DEET protected significantly longer than the controlled-release formulation containing 20 percent DEET (P<0.001).

No non-DEET repellent fully evaluated in this study was able to provide protection that lasted more than 1.5 hours. Only the soybean-oil–based repellent was able to provide protection for a period similar to that of the lowest-concentration DEET product we tested (94.6 and 88.4 minutes, respectively).

The IR3535-based repellent protected against mosquito bites for an average of 22.9 minutes. The citronella-based repellents we tested protected for 20 minutes or less. There was no significant difference in protection time between the slow-release formulation containing 12 percent citronella and the formulation containing 5 percent citronella (P=0.07) or the two formulations containing 10 percent citronella (P=0.16 and P=0.80). The repellent containing only 0.05 percent citronella provided less protection than the Skin-So-Soft mineral-oil–based moisturizer (Avon) (P<0.001). Repellent-impregnated wristbands, containing either 9.5 percent DEET or 25 percent citronella (by weight), protected the wearer for only 12 to 18 seconds, on average.

In arm-in-cage studies, testing must be conducted with insertions at limited intervals, with a new batch of mosquitoes for each test, because continuous exposure may cause mosquitoes to fatigue or may induce prolonged blockage of their antennal chemoreceptors, both of which will prevent further biting. Conducting tests of a repellent in which the arm is inserted into the cage at fixed intervals, however, has some obvious limitations. A repellent might stop working between the removal of the arm and the subsequent insertion, but the failure would not be detected until the next scheduled insertion, causing an inflated measure of the duration of protection provided by that repellent. In our study, the greatest risk of overestimation of complete-protection times would affect the repellents that were tested with once-hourly insertions into the cage. According to our protocol, however, hourly insertions were only used by subjects who found that a repellent initially protected them for more than four hours. Only the two highest-concentration DEET-based repellents in our study (20 percent and 23.8 percent DEET) qualified for once-hourly insertions by some of the subjects, and the range of protection these repellents afforded (180 to 360 minutes) is consistent with previously published reports of the efficacy of DEET. Any rounding errors resulting from the intervals between insertions into the cage would also tend to overestimate the efficacy of the other repellents we tested, and 11 of the 12 non-DEET products still had mean complete-protection times of less than 23 minutes.

After the original studies for this article were completed, a new botanical repellent was introduced in the United States. The repellent contains oil of eucalyptus and is marketed under two names: Repel Lemon Eucalyptus Insect Repellent (WPC Brands) and Fite Bite Plant-Based Insect Repellent (Travel Medicine). We evaluated this type of repellent using the same testing methods in six subjects (five men and one woman). In one subject, a localized cutaneous reaction developed after the first test, and the subject discontinued the study. All other subjects completed three tests each of the repellent. The repellent had a mean (±SD) complete-protection time of 120.1±44.8 minutes, with a range of 60 to 217 minutes.

Discussion

Protection against arthropod bites is best achieved by avoiding infested habitats, wearing protective clothing, and applying insect repellent. The insect repellents that are currently available to consumers are either synthetic chemicals or are derived from plants. The most widely marketed chemical-based insect repellent is DEET, which has been used worldwide since 1957. DEET is a broad-spectrum repellent that is effective against many species of mosquitoes, biting flies, chiggers, fleas, and ticks. The protection provided by DEET is proportional to the logarithm of the dose; higher concentrations of DEET provide longer-lasting protection, but the duration of action tends to plateau at a concentration of about 50 percent. Most commercially available formulations now contain 40 percent DEET or less, and the higher concentrations are most appropriate to use under circumstances in which the biting pressures are intense, the risk of arthropod-transmitted disease is great, or environmental conditions promote the rapid loss of repellent from the surface of the skin.In our study, a formulation containing 23.8 percent DEET provided an average of five hours of complete protection against A. aegypti bites after a single application. Depending on the formulation and concentration tested, DEET-based repellents have been shown in other studies to provide complete protection against arthropod bites for as long as 12 hours, even under harsh climatic conditions.

The most recent addition to the synthetic insect repellents on the market in the United States is IR3535, which is classified by the Environmental Protection Agency as a biopesticide because of its structural similarity to the amino acid alanine. This repellent has been used in Europe for more than 20 years and was approved for use in the United States in 1999. In our tests, this repellent fared poorly, yielding a mean complete-protection time that was one quarter that of the lowest-concentration DEET product we tested (22.9 vs. 88.4 minutes).

Skin-So-Soft Bath Oil, which consumers commonly claim has a repellent effect on insects, provided only a mean of 9.6 minutes of protection against aedes bites in our study. This extremely limited repellent effect has previously been documented in other studies.

Thousands of plants have been tested as potential botanical sources of insect repellent. Most plant-based insect repellents currently on the market contain essential oils from one or more of the following plants: citronella, cedar, eucalyptus, peppermint, lemongrass, geranium, and soybean. Of the products we tested, the soybean-oil–based repellent was able to protect from mosquito bites for about 1.5 hours. All other botanical repellents that we tested in our initial studies, regardless of their active ingredients and formulations, gave very short-lived protection, ranging from a mean of about 3 to 20 minutes. Preliminary studies suggest that the oil-of-eucalyptus products will confer longer-lasting protection than other available plant-based repellents.

Most alternatives to topically applied repellents have proved to be ineffective. No ingested compound, including garlic and thiamine (vitamin B1), has been found to be capable of repelling biting arthropods. Small, wearable devices that emit sounds that are purported to be abhorrent to biting mosquitoes have also been proved to be ineffective. In our study, wristbands impregnated with either DEET or citronella similarly provided no protection from bites, consistent with the known inability of repellents to protect beyond 4 cm from the site of application.

Multiple factors play a part in determining how effective any repellent will be; these factors include the species of the biting organisms and the density of organisms in the immediate surroundings; the user’s age, sex, level of activity, and biochemical attractiveness to biting arthropods; and the ambient temperature, humidity, and wind speed. As a result, a given repellent will not protect all users equally. Examination of the ranges of complete-protection times in Table 1 shows variation in the ability of each repellent to protect different subjects. Thus, these times should be taken not as absolute values but, rather, as an indication of the relative effectiveness of the tested repellent products.

Our study shows that only products containing DEET offer long-lasting protection after a single application. Certain plant-derived repellents may provide short-lived efficacy, which may be sufficient when arthropod bites are primarily a nuisance. Frequent reapplication of these repellents would partially compensate for their short duration of action. However, when one is traveling to an area with prevalent mosquito-borne disease that could be transmitted through a single bite, the use of non-DEET repellents would seem to be ill-advised. Given our findings, we cannot recommend the use of any currently available non-DEET repellent to provide complete protection from arthropod bites for any sustained outdoor activity.

Although this study shows that DEET-based products can be depended on for long-lasting repellent effect, they are not perfect repellents. DEET may be washed off by perspiration or rain, and its efficacy decreases dramatically with rising outdoor temperatures. DEET is also a plasticizer, capable of dissolving watch crystals, the frames of glasses, and certain synthetic fabrics.

Despite the substantial attention paid by the lay press every year to the safety of DEET, this repellent has been subjected to more scientific and toxicologic scrutiny than any other repellent substance. The extensive accumulated toxicologic data on DEET have been reviewed elsewhere. DEET has a remarkable safety profile after 40 years of use and nearly 8 billion human applications. Fewer than 50 cases of serious toxic effects have been documented in the medical literature since 1960, and three quarters of them resolved without sequelae. Many of these cases of toxic effects involved long-term, heavy, frequent, or whole-body application of DEET. No correlation has been found between the concentration of DEET used and the risk of toxic effects. As part of the Reregistration Eligibility Decision on DEET, released in 1998, the Environmental Protection Agency reviewed the accumulated data on the toxicity of DEET and concluded that “normal use of DEET does not present a health concern to the general U.S. population.” When applied with common sense, DEET-based repellents can be expected to provide a safe as well as a long-lasting repellent effect. Until a better repellent becomes available, DEET-based repellents remain the gold standard of protection under circumstances in which it is crucial to be protected against arthropod bites that might transmit disease.

Source Information

From Chapel Hill Dermatology, Chapel Hill, N.C. (M.S.F.); and the Florida Medical Entomology Laboratory, University of Florida, Vero Beach (J.F.D.).

 

 

Introduction
Transpiration is the process through  which water is lost from a plant by evaporation. Water is taken into a plant through roots and root hairs by osmosis, and it exits the plant through ting openings on the underside of leaves known as stomata. Oxygen and carbon dioxide are exchanged through the stomata. Transpiration is also the major mechanism that powers the movement of water throughout a plant. This transportation of water through the plant is due to water potential. Water potential is the potential energy created by the water molecules within the plant stem. Water always flows from areas of high water potential to areas of low water potential. Gravity, pressure, and solute concentration are all factors determining water potential in a plant.

There are three main kinds of cells in plants. the most abundant is parenchyma cells. These cells are mainly unspecialized and make up the mesophyll layer in leaves. Most parenchyma cells store food such as starch to be used later in the plant. Sclerenchyma cells are lignified and dead at maturity. These cells make up fibers and have thick secondary cell walls. They serve as support in plants. Collenchyma cells can be found in young stems and leaves. They are living at maturity and have thick primary cell walls.  There are also three types of tissues found in plants — xylem, phloem, and epidermal. The epidermal cells make up the outermost layer of cells on a plant and function in protecting the plant. Xylem is the water conducting tissue of the plant, while phloem is the food conducting plant tissue.

 

In this experiment, four bean plants will be used to test transpiration rates under different environmental conditions. The conditions included a normal room setting, exposure to a fan, heat lamp, and moist environment ( air misted and plant covered with plastic bag). Data will be obtained from each setting to determine if the various conditions affected the rate of water loss from leaves.

Hypothesis
Under the setting in which the plant is prayed with water and then covered in a plastic bag to create a moist environment, there will be the lowest rate of transpiration.

Materials
9A
Materials used for part A included a graduated cylinder, parafilm, distilled water, bean plant, scalpel, watch, fan, heat lamp, spray bottle, plastic bag to cover plant, and a metric scale.
9B
Materials needed for part B included a microtome, single edge razor blade, paraffin, 50% ethanol, toluidine blue stain, distilled water, 50% glycerine, microscope slide, petri dishes, and compound microscope.

 

Methods
9A
First make a potometer by filling the graduated cylinder with water and covering it securely with parafilm. Poke a hole in the parafilm. Remove the root from the rest of the plant and insert the plant into the parafilm hole so that the end of the stem is below the water level in the graduated cylinder. Record the initial water level in the potometer. Weigh the potometer with the plant and record the initial mass.  Expose this plant to one of the four conditions (misted plant), and take readings of the potometer mass every 10 minutes for a total of 30 minutes. Record this data in your data table.

9B
In this part of the experiment, a cross-section of a leaf will be observed. Cut the stem of a non-woody plant about 5mm longer than the depth of the microtome. Hold the stem vertically in the microtome and pour melted paraffin around it. Allow the paraffin to cool and harden around the stem. Use a razor blade to cut off the excess stem above the paraffin. Slightly turn the microtome to expose a thin layer of the stem. Slice several thin layers of the stem from the microtome and place these slices in a petri dish containing 50% ethanol for 5 minutes. Move the slices to another petri dish containing toluidine blue stain for  1-2 minutes. Rinse the slices and then mount each section on a microscope slide in a drop of 50% glycerine. Add a cover slip and observe under a compound microscope. Draw the stem cross-section.

 

Calculating Leaf Surface Area

 

Data 9A

Transpiration Rate

 

Transpiration Rate

Data 9A

Leaf Cross Section

 

Questions

1. Calculate the average rate of water loss per minute for each of the following treatments:
Room:    0 ml/min
Fan:        .53 ml/min
Light:      .367 ml/min
Mist:       .23 ml/min

2.

4. A plant with its stomata closed prevents water that is needed by the plant from escaping.

5. Some plants, such as CAM plants, have adaptations to prevent water loss. These plants have their stomata closed during the day (hottest part of the day when water loss would be greatest) and their stomata open during the night when its cooler to carry out photosynthetic reactions. This reduces water loss from leaves.

Error Analysis
During this experiment, there were many complications that arose. When using potometers, sealing the potometers was difficult affecting  the rate of water loss. By changing the procedure and massing the graduated cylinders at timed intervals, more accurate data was obtained.

Conclusion
Although the misted plant did have a low rate of water loss, it was not the lowest transpiration rate observed.  The lowest transpiration rate came from the plant at room temperature, the control plant. The plants exposed  to the heat lamp and fan showed the highest rate of water loss as expected.

* Art work from Lab Bench    Http://www.biology.com

 

 

Introduction

Most of the water a plant absorbs is not used for a plant’s daily functioning. It is instead lost through transpiration, the evaporation of water through the leaf surface and stomata, and through guttation, which is the loss of water from the vascular tissues in the margins of leaves.

There are three levels of transport in plants: uptake and release of water and solutes by individual cells, short distance cell to cell transport at tissue and organ levels, and long distance transport of sap by xylem and phloem at the whole plant level. The transport of water is controlled by water potential. Water will always move from an area of high water potential to an area with low water potential. This water potential is affected by pressure, gravity, and solute concentration.

Water moves into the plant through osmosis and creates a hydrostatic root pressure that forces the water upward for a short distance, however, the main force in moving water is the upward pull due to transpiration. This pull is increased by water’s natural properties such as adhesion and cohesion. Transpiration decreases the water potential in the stele causing water to move in and pull upward into the leaves and other areas of low water potential. Pressure begins to build in the leaves, so to prevent downward movement, guttation occurs. Guttation occurs through leaf openings on the leaf margins called hydrathodes. Loss of water through transpiration can be facilitated by the opening and closing of the stomata depending on environmental conditions.

There are three types of cells in plants: parenchyma, sclerenchyma, and collenchyma. Parenchyma cells are the most abundant and are not specialized. They are found in the mesophyll of leaves, the flesh of fruits, the pith of stems, and the root and stem cortex. Sclerenchyma are elongated cells that make up fibers. They have thick secondary walls and the protoplasts often die as they grow older. They are used for support and are found in vascular tissue. Collenchyma cells are living at maturity and have a thickened secondary wall.

 

Hypothesis

 

In Lab 9A, all of the plants in this experiment will lose water through transpiration, but those affected by the heat sink and the fan will lose a larger amount of water due to the environmental conditions. This transpiration will pull water from the potometer into the plant. The structure and cell types of a stem cross-section can be observed under a microscope.

 

Materials

 

Exercise 9A: Transpiration

The materials needed for this exercise were a pan of water, timer, a beaker containing water (heat sink), scissors, 1-mL pipette, a plant cutting, ring stand, clamps, clear plastic tubing, petroleum jelly, a fan, lamp, spray bottle, a scale, calculator, and a plastic bag.

Exercise 9B: Structure of the Stem

The materials needed for this exercise were a nut-and-bolt microtome, single-edge razor blade, plant stems, paraffin, 50% ethanol, distilled water, 50% glycerin, toluidine blue O stain, a microscope slide and cover slip, pencil, paper, and a light microscope.

Methods

Exercise 9A: Transpiration

The tip of the pipette was placed in the plastic tubing and they were submerged in a tray of water. Water was drawn into the pipette and tubing until no bubbles were left. The plant stem was cut underwater and inserted into the plastic tubing. Petroleum jelly was immediately placed around the tube edging to form an airtight seal around the stem. The tubing was bent into a “U” shape and two clamps were used on the ring stand to hold the potometer in place. The potometer was allowed to equilibrate for ten minutes.

The plant was exposed to a fan, which was placed one meter away and set on low speed. The time zero reading was recorded and then it was continually recorded every three minutes for 30 minutes. After the experiment, all the leaves were cut off the plant and massed by cutting a one cm2 box and massing it.

Exercise 9B: Structure of the Stem

A nut-and-bolt microtome was obtained and a small cup was formed by unscrewing the bolt. The stem was placed in the microtome and melted paraffin was poured around the stem. The paraffin was allowed to dry and the excess stem was cut off. The bolt was twisted just a little and then cut with the blade. The slice was placed in the 50% ethanol. The slices were left in the ethanol for five minutes. Using the forceps, the slices were moved to a dish of the toluidine blue O stain and left for one minute. The sections were rinsed in distilled water. The section was mounted on the slide with a drop of 50% glycerin. A cover slip was placed over the slide. The cross section was observed under a light microscope and drawn.

 

Results

Table 9.1: Individual Potometer Readings

 

Time (min)	Beginning (0)	3	6	9	12	15	18	21	24	27	30
Reading (mL)	.02	.03	.04	.05	.06	.07	.09	.10	.11	.13	.13

 

Class Potometer Readings

 

Time (min)	Beginning (0)	3	6	9	12	15	18	21	24	27	30
Room	.53	.54	.55	.56	.57	.58	.59	.60	.61	.62	.63
Mist	.34	.36	.38	.40	.42	.43	.43	.44	.45	.45	.46
Light	.67	.68	.69	.70	.71	.72	.73	.74	.75	.77	.79
Fan	.02	.03	.04	.05	.06	.07	.09	.10	.11	.13	.13

 

Mass of leaves = 1.1 g
Leaf Surface Area = 0.0044 m2

 

Table 9.2: Individual Water Loss in mL/m2

 

Time Interval (minutes)
	0-3	3-6	6-9	9-12	12-15	15-18	18-21	21-24	24-27	27-30
Water Loss (mL)	.01	.01	.01	.01	.01	.02	.01	.01	.01	0
Water Loss per m2	2.27	2.27	2.27	2.27	2.27	4.55	2.27	2.27	2.27	0

 

 

 

Table 9.3: Class Average Cumulative Water Loss in mL/m2

 

	Time (minutes)
Treatment	0	3	6	9	12	15	18	21	24	27	30
Room	0	5	5	5	5	5	5	5	5	5	5
Light	0	2.5	2.5	2.5	2.5	2.5	2.5	2.5	2.5	4	4
Fan	0	2.27	2.27	2.27	2.27	2.27	4.55	2.27	2.27	2.27	0
Mist	0	4.17	4.17	4.17	4.17	2.08	0	2.08	2.08	0	2.08

 

 

Analysis of Results

Calculate the average rate of water loss per minute for each of the treatments:
Room: 1.67 mL/m2
Fan: 0.76 mL/m2
Light: 0.93 mL/m2
Mist: 0.83 mL/m2

 

Explain why each of the conditions cause an increase or decrease in transpiration compared with the control.

 

 

How did each condition affect the gradient of the water potential from stem to leaf in the experimental plant?

 

The light and the fan decreased the water potential in the leaves and water moved up the stem by transpiration pull. The room temperature had little or no effect on the water potential. The mist increased the water potential of the air causing less transpiration to occur from the leaves.

 

What is the advantage to a plant of closed stomata where water is in short supply? What are the disadvantages?

 

The closing of the stomata would prevent transpiration of water and minimize this loss if water was in short supply. It is a conservational adaptation. However, closing stomata prevents the exchange of gases in plants and limits their carbon supplies.

 

Describe several adaptations that enable plants to reduce water loss from their leaves. Include both structural and psychological adaptations.

 

Plants that are adapted to drier climates are called xerophytes. Some of these plants have adapted small, thick leaves with a reduced surface area. They may also have a thickened cuticle to protect themselves from the environment. The stomata may be sunken into pits. Some xerophytes shed their leaves during the driest seasons and others can store water such as cacti. CAM plants uptake CO2 at night and change it into crassulacean acid that can be broken down during the day for sugars. These plants can close their stomata during the day.

 

Why did you need to calculate leaf surface area in tabulating your results?

 

The surface area has to be calculated because this greatly affects the amount of water lost through transpiration. Smaller leaves may lose less water than the larger ones, but by calculated water loss by surface area creates comparable data that is constant and consistent.

 

Error Analysis

 

This lab had many opportunities for error. The potometer set up was a complicated procedure. If any air bubbles were present in the plastic tubing, it could cause drastic error to occur. Any miscalculations or inaccurate weighing could also account for error.

 

Discussion and Conclusion

 

Transpiration in plants is controlled by water potential. This change in water potential in leaves causes a gradient by which water can be moved upward. When the water potential of the air was increased by the mist and plastic bag, less water evaporated from the leaves, decreasing the water potential gradient between the root and stem. This decreased the transpiration pull. The fan and floodlight simulated environmental conditions such as wind, heat, and intense light. These conditions increase the amount of water transpired by plants. This in turn increased the water potential gradient causing more water to be pulled through the stem. The control plant should have had normal rates of transpiration.

The stem must have specialized cells for support and transport. The epidermis is the outermost layer of the stem. The xylem is a transport tube for water, and the phloem transports food and minerals through the plant. Parenchyma are non-specialized cells and are located in the interior. The tougher sclerenchyma and collenchyma make up the structural outer support of the epidermis and the transport tubes of phloem and xylem.

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Arkansas is essentially a forest state because more than half of the state is covered with trees.  The climate and soils of Arkansas also support a great variety of trees, both conifers and deciduous.  Trees are one of Arkansas’ most important crops.  Forests are also valuable in preventing erosion, in offering parks and recreational areas, and in providing homes for wildlife.  In addition, many trees have been introduced into the state as ornamentals.

Leaf collecting is a good way to learn the trees native to your area.  Collecting leaves will also help you to learn leaf margins, shapes, and  venations and how to use different taxonomic keys to identify trees.

Materials needed:

• leaf press
• black ink pen
• pencil
• small notebook to record leaves collected
• scissors
• Elmer’s glue
• art paper, poster board, etc. for mounting
• labels
• taxonomic keys (Trees of Arkansas published by the Arkansas Forestry Commission)

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Directions for making a leaf press:
1. Cut 10-15 pieces of corrugated cardboard 30 cm by 50 cm in size.
2. Cut several sheets of newspaper the same size as the cardboard.
3. Lay sheets of newspaper between each cardboard layer sandwich style.
4. A piece of wood may be added to the top and bottom to better “press” the leaves flat
5. Use two stretch belts or cords to bind the press together.
6. Leave the press in an area so that air can circulate &  more quickly dry the leaves.

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Getting started with your collection:
1. Study the shapes, margins, venations, tips, bases, etc. in your Trees of Arkansas book.

Click here to view reference page

2. Learn to distinguish simple leaves (one blade) from compound leaves (multiple leaflets) and conifers (evergreens) from deciduous (lose leaves) trees.
3. Learn to distinguish a tree from a shrub. (Trees with a single trunk)
4. leaves attach to twigs at NODES. INTERNODES are the distance between leaves on a twig.
5. Gather your collecting materials together – press, pencil, scissors, & small notebook.
6. Always get permission before collecting leaves on someone else’s property.
7. Be sure to collect at least two of each type of leaf so both the bottom & top side of the leaf can be shown in your collection.
8. Place leaves in your press immediately after collecting them so they do not start to dry out and wrinkle.
9. Record the name of each leaf, date collected, and place collected in your notebook as you collect.
Also record tree characteristics such as shape of the crown, color and type of bark, etc.

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Collecting:
1. Remember to collect two of every type of leaf!
2. Carefully remove an entire leaf, not a leaflet, from the tree, and place this in your press between newspaper layers.
3. If leaves are damaged or torn, don’t use them because you will not receive credit.
4. Make sure that none of the leaf parts extend beyond the edge of the press.
5. You may also collect & press seeds and/or fruits from some trees if they fit in your press.
6. Leave the leaf in the press for 3 – 5 days depending on its thickness and moisture content.  .
7. Keep the press in an area where air is circulating (in front of a fan).

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Labeling and identifying:
1. Obtain printed labels from your teacher.
2. Use only black ink to write labels, & do not mark out or white out mistakes on the labels; rewrite them.
3. Use taxonomic keys to identify each leaf, and include both the scientific & common name of the tree on the label.
4. Determine the shape, margin, tip, base, and venation of your leaf and whether it is a simple or compound leaf; record this on your label.
5. Use you key to give a description of the tree, not the leaf.
6. Research uses for the tree, its fruit, etc. and record on your label.
7. Tell if the leaf is deciduous or coniferous.

Click here for more labels

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Mounting leaves: 

Paste Label Here             Pg#

LEAF COLLECTION Name Date Period Subject/Teacher

1. Use pieces of cut poster board or art paper to mount your leaves.  Make sure all sheets are uniform in size! (The size of your sheets will be determined by your largest leaf.)
2. Use Elmer’s glue to adhere two leaves to each page — one showing the upper surface of the leaf and the other showing the underside of the leaf.
3. Each page should have only one type of leaf on it.
4. Arrange the leaves so they do not overlap each other and so there is room to glue the label in the lower right hand corner.  The leaves should look nice on the page.
5. On compound leaves, mount the topside of the complete leaf and then mount the underside of a single leaflet. Make sure the leaflet comes from another leaf to receive credit!
6. Use a small amount of Elmer’s glue to adhere the completed label in the lower right hand corner of the page.
7. LET THE PAGES DRY COMPLETELY BEFORE ASSEMBLING THEM TOGETHER IN YOUR COLLECTION OR THE PAGES WILL STICK TOGETHER!!!!!
8. Once the pages are dry, lay them in the correct order (see your list of required leaves), and then number the pages in the lower right corner with black ink.
9. Make a stiff front and back cover for your collection from poster board, cardboard, wood, etc.  Include the following items on your cover:

• title
• your complete name
• date collection turned into teacher
• class period

10. Use ribbon, string, etc. to bind the pages together or assemble the collection in a scrapbook or art book.  DO NOT COVER THE LEAVES WITH PLASTIC!!!

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Required leaves:
1. Only native, Arkansas trees may be used.  Refer to your Trees of Arkansas book.
2. Leaves must be in perfect condition without damage or tears.
3. No more then 4 oaks are allowed in the collection.
4. No fruit trees such as apple, pear, orange, peach, etc. are allowed.
5. Place the following leaves in your collection first and in this order:

1. sweet gum
2. American sycamore
3. pine (any type)
4. flowering dogwood
5. redbud
6. ash (any type)
7. redbud
8. Eastern red cedar
9. maple
10. willow
11. pecan
12. pin oak
13. willow oak
14. water oak
15. elm (any type)

 16 – 20  Any other Native Arkansas leaves

6. The remaining leaves that you include must be trees native to Arkansas!

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*Pre AP Biology is required to collect 30 leaves including the 15 required.

 *Biology I is required to collect 20 leaves including the 15 required.