The Comparison of Responses of the Same Level of Exercise on the Heart Rate and Blood Pressure of Boys and Girls

By: Joyce Acen

Introduction:

Question:

Will the girls’ and boys’ responses to the same level of exercise differ on its effect to their heart rate and blood pressure?

Background:

The functions of the human cardiopulmonary physiology can be broken down into the circulatory system and the respiratory system. The main goal of these two systems is to maintain homeostasis. Homeostasis can be described as a type of condition where the internal continuity of an individual has to keep steadiness, regardless of any external changes from the outside environment. (Biozone, 252) External changes from the outside environment may include factors such as excitement, stress, exercise, diet, and many more. Exercise can be carried out because the human body can endure a high level of exercise during a prolonged period of time. (Biozone, 226) In this experiment, we will focus on how exercise, in particular, affects the cardiopulmonary physiology of girls and boys.

Exercise places strenuous activity on the human body. When an individual exercises, the blood flow must level up to the demands being made on the individual’s muscles, heart and lungs. (Biozone, 225) Blood flow increases so that the blood does not clot and so that it can supply oxygen and all of the necessary nutrients to the tissues and organs. An essential reason to why oxygen and nutrients are needed is because they are the ingredients that allow the heart to continue pumping blood throughout the venous, arterial, and portal systems. (Biozone, 225) The heart is one of the main organs involved in the human cardiopulmonary physiology because it keeps blood moving through the contraction of its valves. More specifically, the right side of the heart pumps blood to the lungs so that it can get oxygenated, while the left side of the heart pumps blood to the body.

When an individual exercises, the generated heat must be dissipated, oxygen demands must increase, and waste products must be produced. If we imagine that an individual has begun exercising, the oxygen uptake increases because there is a higher demand for energy. Oxygen is the ultimate source of energy that allows ATP to be generated. More ATP must be made in order for homeostasis to be maintained. After a prolonged period of exercise, lactic acid accumulation begins to occur in the tissues of the body because energy for oxygen uptake can no longer be supplied after a certain period of time of exercise. (Biozone, 308) When the individual ends exercising, it takes time for the oxygen uptake to return to its resting level because the lactic acid that had accumulated in the tissues must be broken down into CO2 and H2O. The individual breathes deeply during this time because heavy breaths of oxygen must be taken in for the lactic acid to be broken down.

A process which allows human beings to take in oxygen and remove carbon dioxide would be gas exchange. The set of mechanisms within this process allow cellular respiration to occur. Cellular respiration results in the release of energy that comes from organic compounds. These organic compounds generate waste materials that must be removed through exhalation. (Biozone, 191)

Hypothesis:

Since the human body is put under strenuous physical activity during exercise, it will require faster circulation and respiration throughout the body. Because one of the factors that affect the cardiopulmonary function is gender, if we test girls and boys under the same level exercise, then the boys will have higher blood pressure and a higher heart rate and will recover faster because they can supply their bodies with oxygen at a quicker rate than girls.

 

Overview of Experiment:

Through this lab, the girls’ and boys’ responses to the same level of exercise will be observed and compared to find similarities and differences in its effects to their heart rate and blood pressure. The hypothesis will be tested through the method of having measured the individuals’ blood pressure with the sphygmomanometer.

Materials and Methods:

            The scientist must have his/her breathing rates measured under different conditions with the use of a stopwatch, and a sphygmomanometer. To begin experimentation, the individual being tested must have his/her resting blood pressure measured. To do this, the sphygmomanometer must be placed above the elbow so that the wire connected to the monitor and the pressure cuff goes over the brachial artery. The cuff should be inflated to about 150mm Hg each time a measurement it taken. Once the resting blood pressure has been found, the individual being tested should begin exercising for 1 minute without a break. Immediately after exercise, the individual should have his blood pressure measured. Then every 2 minutes after the individual has stopped exercising, the blood pressure should also be measured until a total of 10 minutes has passed.

Results:

The overall results showed that the boys had higher systolic and diastolic pressures, while their heart rate pulses were not significantly different from the data results of the girls’ heart rate pulses.

The results for the boys indicated that their initial systolic pressure was at 121 mm Hg, their diastolic pressure was at 76 mm Hg, and their heart rate pulse was at 75 beats per minute (bpm). Their highest systolic pressure was 146 mm Hg at 0 minutes, 89 mm Hg for the diastolic pressure at 0 minutes, and 119 bpm for the pulse at 0 minutes. 0 minutes represents the immediate time after exercise was finished. The boys’ lowest systolic pressure was 121mm Hg at 6 minutes, 70 mm Hg for the diastolic pressure at 8 minutes, and 71 bpm for the pulse at 4 minutes. The recovery rate was measured by finding the difference between immediate exercise and 2 minutes. The boys’ systolic pressure recovery rate decreased by 11 mm Hg, their diastolic pressure decreased by 2 mm Hg, and their pulse rate decreased by 43bpm.

The results for the girls showed that their initial systolic pressure was 104 mm Hg, 67 mm Hg for their diastolic pressure, and 72 bpm for their pulse rate. Their highest recordings came in at 0 minutes, immediately after exercise. They were 128 mm Hg for the systolic pressure, 82 mm Hg for the diastolic pressure, and 120bpm for the pulse rate. Their lowest data points were consistent with the basal time because the systolic pressure was 104 mm Hg at their basal time (before exercise), their lowest diastolic pressure was 67 mm Hg at their basal time, and their lowest pulse time was 72bpm at the basal time. The girls’ systolic pressure recovery rate was decreased by 7 mm Hg, their diastolic pressure recovery rate was decreased by 6 mm Hg, and their pulse rate decreased by 39bpm.

boys and girls heart rate

 

This table summarizes the averages of systolic pressure, diastolic pressure, and the heart rate pulses for both genders.

Discussion:

The hypothesis for this laboratory report stated that if we test girls and boys under the same level exercise, then the boys will have higher blood pressure and a higher heart rate and will recover faster because they can supply their bodies with oxygen at a quicker rate than girls.

Looking at the data results, the boys did have higher systolic and diastolic pressures than the girls, but their pulse recordings were fairly similar to that of the girls’ pulse data records. The boys’ systolic pressure was higher by about 20 mm Hg throughout each recording. The boy’s diastolic pressure was higher by about 5 to 10 mm Hg than that of girls.  But both genders’ pulse rates were ranged from 70bpm to 120bpm. As for the recovery rate, the data collected indicates that the difference rate after immediate exercise to 2 minutes was higher for the boys than for the girls because the decrease from the recordings after immediate exercise to 2 minutes was more extreme in mm Hg and in heart beats per minute for the boys. So overall, the data collected supports the hypothesis for the most part except for the statement made about the heart rate being higher for boys than for the girls. The hypothesis for the heart rate difference was incorrect because the assumption about the relationship between systolic and diastolic pressure was correlated to the heart rate pulse recordings.

One source stated that the effect of exercise on blood pressure will vary depending on factors such as gender (Sheryl R, 2010). Therefore, the results for the heart pulse rate conflict with the hypothesis because theoretically the heart rates for both genders should be somewhat different.

Some weaknesses identified within the experimental set would be inaccurate timing, and equal intensities of exercise among all students. Inaccurate timing seemed to be an issue because it was difficult to gather recordings exactly 2 minutes from every previous recording. This was a problem because the sphygmomanometer would sometimes automatically shut down causing a prolonged timing error between recordings. The intensity of exercise was not the same among all students which could have contributed to inaccurate data. Some students exercised harder for the 1st minute than others which made their systolic and diastolic pressures significantly higher.

Literature Cited:

Brainbridge-Smith, Lissa, Richard Allan, and Tracey Greenwood. “Energy and Exercise.” Print. Rpt. in Senior Biology 2 2009. 8th ed. BIOZONE International, 2008. 307-08. Print.

Brainbridge-Smith, Lissa, Tracey Greenwood, and Richard Allan. “Exercise and Blood Flow.” Print. Rpt. in Senior Biology 2 2009. 8th ed. BIOZONE International, 2008. 225-26. Print.

Greenwood, Tracey, Lissa Brainbridge-Smith, and Richard Allan. “Introduction to Gas Exchange.” Print. Rpt. in Senior Biology 2 2009. 8th ed. BIOZONE International, 2008. 191. Print.

R, Sheryl. “The Relationship Between Diastolic Blood Pressure and Exercise.” Find Health, Education, Science & Technology Articles, Reviews, How-To and Tech Tips At Bright Hub – Apply To Be A Writer Today! 3 May 2010. Web. 19 Feb. 2011. <http://www.brighthub.com/health/fitness/articles/41153.aspx>.

 

Biochemical Activities

Biochemical Activities
By: Amanda Goldner

Table of Contents

Lab A: Media Preparation
Lab 1: Inoculations, Smear Preps, and Simple Staining
Lab 2: Gram Staining and Using the Spectrophotometer
Lab 3: Negative, Acid Fast, Endospore, and Capsule Staining
Lab 4: Biochemical Activities of Bacteria I ……………………………………….… 3

Lab 5: Bergey’s Manual (Practical) …………………………………………………  11
Lab 6: Biochemical Activities of Bacteria II, Unknown Identification, and
Rapid Multitests …………………………………………………………………………………………… 13
Lab 7: Isolation of Microorganisms From Soil
Lab 8: Eukaryotes I: Fungi
Lab 9: Eukaryotes II: Algae and Protozoa
Lab 10: Poster Presentations

Lab 4: Biochemical Activities of Bacteria I
I. Purpose

The objective of this lab was to run differential tests on Escherichia coli, Pseudomonas fluorescens, Bacillus subtilis, and Proteus vulgaris. This was done to determine such characteristics as ability to ferment lactose/glucose/sucrose, produce hydrogen sulfide/acids/acetoin, and utilize urea or citrate.

II. Materials
The materials needed for this lab were as follows:

– inoculating loop
– disposable gloves
– inoculating needle
– permanent marker
– Bunsen burner
– 1 Eosin Methylene Blue (EMB) agar plate
– 1 Hektoen agar plate
– 1 McConkey agar plate
– Barritts A reagent
– Barritts B reagent
– Kovac’s reagent
– biohazard container
– 4 Pasteur pipettes
– 4 Sulfide-Indole Motility (SIM) agar deeps
– 4 Triple Sugar Iron (TSI) agar slants
– 4 urea agar slants
– 4 Simmons Citrate Agar (SCA) slants
– 4 Glucose Phosphate Broth tubes
– 4 empty test tubes
– tryptic soy agar plate cultures (pure) of:

~ Escherichia coli

~ Bacillus subtilis

~ Pseudomonas fluorescens
~ Proteus vulgaris

III. Methods

McConkey, EMB, and Hektoen Agar Plates

    The McConkey, Eosin Methylene Blue (EMB), and Hektoen plates were each inoculated with a streak of every bacterial variety listed in the Materials section of this report. Each plate had four resulting one-inch streaks, one in each quadrant and each made up of a different species inoculum. Aseptic technique was maintained in all bacterial transfers (ex:  flaming the inoculating loop at proper times).  The plates were labeled on the bottom with permanent marker, split into four quadrants for each streak.  The genus species initials for each bacterial type were written down in the respective quadrant in which that species was located.  All three inoculated plates were then placed on a storage rack to be incubated for 24 hours.

Triple Sugar Iron (TSI) Agar Slants

    The four Triple Sugar Iron agar slants were each inoculated via inoculating needle with a different species of bacteria (those listed in Materials section).  Proper aseptic technique concerning inoculation and test tubes was maintained in all bacterial transferals.  The TSI slant was first stabbed with the needle, before the needle was dragged in a zigzag pattern across the surface of the slant from the lower-elevation end to the higher-elevation end.  The needle was not flamed between the stabbing and streaking of the agar.  Once this procedure had been applied to all slants, the tubes were placed in the storage rack for a 24-hour period incubation.

Simmons Citrate (SCA) and Urease Agar Slants

    One Simmons Citrate Agar slant and one urease slant was reserved for each species of bacteria listed in the Materials section above.  All slants were inoculated via inoculation loop in a zigzag pattern across the surface of the slant from the lower-elevation end to the higher-elevation end, like the streak configuration seen in the TSI slants.  Aseptic technique pertaining to test tubes and inoculating instruments was maintained in all bacterial transfers.  However, SCA and urease slants were inoculated with a loop and were not stabbed in the butt.  The freshly-inoculated SCA and urease slants were placed to the side in the storage rack for 24 hours of incubation.

Sulfide-Indole Motility Deeps

    The four Simmons Indole Motility agar deeps were inoculated with a vertical stab from an inoculation needle, each with a different type of bacteria (as listed in the Materials section).  Aseptic technique (such as flaming the inoculating needle at proper times) was maintained in all bacterial transferals.  These new culture deeps were then transferred to the storage rack with the rest of the cultures made previously in the lab to be incubated for 24 hours.

Methyl Red and Voges-Proskauer Broth Preparation

    An inoculating loop was used to transfer an inoculum of each species in the Materials section to its own sterile glucose phosphate broth tube, with aseptic technique being strictly followed regarding tube and loop sterilization.  These broth cultures would later be differentiated into eight MR-VP broths, which would then have MR and VP tests run on them using Barritts A and/or B reagent and methyl red indicator.  The cultures were placed to the side for the moment (in the storage rack) to go through a 24-hour incubation period.

———————————————— COMEBACKS ————————————————-

 

Methyl Red and Voges-Proskauer Tests

    The glucose phosphate broth cultures were removed from incubation; half of each tube’s contents was transferred to a new test tube.  To one 4-species tube set, 4-5 drops of methyl red indicator were added and the test tube was shaken.  These tubes were labeled MR and set aside for a few minutes for observation.  To the other four tubes, 10 drops of Barritts A followed by 10 drops of Barritts B were added.  This set of tubes was shaken every three minutes for 15-20 minutes.  Positive results of the methyl red test indicate a pH of 4; the broths developed a red hue.  Negative results of the MR test turned the indicator yellow; this meant the broth cultures had a pH of 6.  As for the Voges-Proskauer test, positive results (red coloration) indicate the presence of acetoin, while negative results involve absence of red tinge and acetoin.

All result conclusions were denoted onto properly-labeled tables found in the Results section of this report.  These outcomes were then studied (see Discussion) to see if the tests had results consistent with the literature values.

Interpreting Results From Plates

    The McConkey agar plates in this experiment were differential, meaning they discerned between bacteria with dissimilarities in characteristics.  Specifically, McConkey differentiated between lactose fermenters and those bacteria that did not ferment lactose.  Bacteria on McConkey agar had lactose positive results if its colonies turned red or pink after the period of incubation.  Lactose negative results mean white, clear, or golden brown dark-centered colonies.
The Eosin Methylene Blue agar plates used in lab were also differential, determining whether a bacterial species can ferment lactose or not.  Positive results were shown by blue-black coloration (E. coli) or pink coloration (Enterobacter).  Since Enterobacter was not used in this experiment, any resultant pink hues mean nothing, and were classified as lactose negative along with lack of colonial coloration.
The third and final type of plate used in this lab was also categorized as differential, but determined differences in pH instead of fermentation ability.  These “Hektoen” agar plates were originally green in color, and changes in pH effect a striking alteration in color:  orange.  Positive lactose fermentation was indicated by orange colonies and/or medium.  No color change of colonies and medium suggested negative lactose fermentation results.  Hektoen plates also revealed hydrogen sulfide production; if this occurred, colonies would have black centers.  Bacteria without black centers were grouped as H2S negative.
All results were recorded into the corresponding tables in the Results section of this report.  These data were later examined to see if the outcomes were similar to those found in previous experiments.

Interpreting Results From TSI Slants

    The area of the tube referred to as the slant encompassed the surface and a few centimeters below, and was inoculated by the needle streak pattern.  The region termed the “butt” comprised the rest of the slant, and was inoculated by the needle stab line.  TSI slants were initially red in color, due to alkaline pH.  If either the slant or the butt changed to a yellow color, it was an acid positive result.  Alkaline positive outcomes stayed red in color.  As can be assumed, acid negative results meant alkaline positive and vice versa. These acid and alkaline results were separate for each region (slant/butt).  If bubbles or cracks were apparent in the agar, then the culture tested gas positive; absence of bubbles/cracks was considered gas negative.  Presence of black material produced H2S positive results, and absence of said black material meant H2S negative results.

All results were logged in table form in the Results section.  Later analysis, as seen in the Discussion section of this report, told if the tests had literature-consistent outcomes.

Interpreting Results From SCA and Urease Slants

    Simmons citrate agar slants fell into the differential category of media.  Inoculation of these slants with a bacterium established whether that species could utilize citrate or not.  A citrate positive outcome involved a change from the original green color to blue.  Citrate negative results did not affect the color, and remained green.  Presence of growth was also noted for this test, with growth positive implying that growth occurred and growth negative meaning lack of growth.
Urease slants also involved determining growth occurrence, with growth positive and negative results implying the same as those for SCA slants.  Pink coloration signified a positive urease test result (the bacteria utilized urea), while negative results (the bacteria did not utilize urea) lacked deviation from original color.

All outcomes were recorded into the corresponding tables in the Results section for later scrutiny, to see if the tests had results consistent with the literature values.

 

 

Interpreting Results From Sulfide-Indole Motility Deeps

    Seven to ten drops of Kovac’s reagent were added to the four SIM agar deeps after removal from incubation.  Within two minutes, results were observed as follows.  Red coloration on the surface of the tube signified indole positive results; absence of said red hue represented an indole negative end result.
All outcomes were detailed into designated tables (see the Results section).  These data were later inspected to see if the results were correct according to literature values.

IV. Results

 

Broths à

 

Methyl Red

 Voges-Proskauer

growth

acid

growth

acetoin

P. vulgaris

turbid throughout until just below the surface

+

+

E. coli

sedimented

at bottom

+

+

P. fluorescens

layered below surface, none beneath center

+

+

B. subtilis

turbid throughout

+

+

Plates

Hektoen Enteric Agar (HE) Eosin Methylene Blue Agar (EMB) McConkey Agar (MAC)

growth

colony color

lactose fermentation

H2S formation

growth

colony color

lactose fermentation

growth

colony color

lactose fermentation

P. vulgaris

+

orange

+

+

+

E. coli

+

orange

+

+

blue

+

+

red

+

P. fluorescens

+

green

+

+

B. subtilis

+

pink

+

+

red

+

 

Slants

Triple Sugar Iron Agar (TSI)

 

Urease Agar

 

Simmons Citrate Agar (SCA)

Butt

Slant

acid

alkaline

gas

H2S production

acid

alkaline

growth

urea hydrolysis

growth

citrate utilization

P. vulgaris

+

+

+

+

+

+

+

E. coli

+

+

+

+

+

P. fluorescens

+

+

+

+

+

+

B. subtilis

+

+

+

+

 

 

  Deeps

Sulfide-Indole Motility Agar

growth

motility

H2S production

indole

P. vulgaris

+

+

E. coli

+

+

+

P. fluorescens

+

B. subtilis

+

+

V. Discussion
This microbiology lab involved the inoculating of many differential media, including:  EMB plates, MR broths, SCA slants, McConkey plates, TSI slants, SIM deeps, Hektoen plates, urease slants, and VP broths.  Each media had a distinct purpose, some even tested for more than one characteristic.  Results were observed after 24 hours of incubation and are interpreted in the following paragraphs along with explanations of the workings of each media type used in this lab.  Every bacterial species in every type of media test had growth present, except for B. subtilis on Hektoen Enteric Agar, as was discussed later on.
Two words used to describe media in this experiment were “differential” and “selective.” Differential media made a distinction between at least two groups of microorganisms.  Differential media made it possible to narrow down the list of possible identities of an unknown bacterial species by biological characteristics alone.  Meanwhile, selective media discriminated against certain types of microbes and favored others.  These media could have been used to make certain that a bacterial colony was part of a particular group (such as gram positive or gram negative), though this function was not taken advantage of.  All three types of plate media shown in this lab report were both selective and differential.
In this experiment, Eosin Methylene Blue (EMB) agar was used to select for gram negative bacteria, inhibiting the growth of any species that were gram positive.  It also determined whether colonies underwent lactose fermentation.  These lactose fermenters typically changed to a blue or black color, because acid production lowered the pH (increasing methylene blue absorption).  Bacteria under the genus Enterobacter were exceptions, and developed a pink hue after incubation on EMB agar.  Non-fermenting bacteria remained colorless.  EMB agar also allowed for recognition of coliform enteric bacteria.  In this experiment, E. coli and B. subtilis acquired the respective colors indicative of lactose fermentation for their genus (on EMB agar), and thus tested lactose positive.  P. fluorescens and P. vulgaris tested negative for lactose fermentation by appearing colorless, however.
McConkey agar was determined to be selective because it prevented the growth of gram positive bacteria by means of bile salts and crystal violet dye.  The pink coloration of the agar came from the crystal violet.  McConkey plates also included lactose, which was converted to lactic acid by lactose fermentation.  This accumulation of acid effected a change in pH (indicated by Neutral Red), and any colonies that ferment acquired a dark pink-red coloration.  Non-fermenters remain opaque and colorless.  MacConkey plates were also differential because of the ability to detect enteric pathogens and coliforms.  In this lab, E. coli and B. subtilis both has positive results, developing a red coloration that revealed their true nature as lactose fermenters.  P. vulgaris and P. fluorescens remained colorless and were thus not labeled lactose fermenters.
Hektoen Enteric agar (HE, HEM, HEK, or HEA) was the only one out of the three agar plates used in this experiment that could sense hydrogen sulfide (H2S) production; colonies that produced H2S took on a black pigmentation.  The agar itself was comprised of lactose, salicin, and bile, which together gave the medium its selective and differential properties.  Coliform enteric bacteria could be identified by using Hektoen media; these turned orange-pink after 24-48 hours.  Salmonella and Shigella typically became blue-green.  HE, like McConkey and EMB, could also distinguish lactose fermenters and non-lactose-fermenters; the region around the fermenting colonies converted to orange.  Results for lactose fermentation of P. vulgaris came out incorrectly in this experiment; the bacteria appeared to be lactose positive but in reality was lactose negative (this error was probably due to wrongful observations or a mix-up).  B. subtilis did not appear to show any growth at all and thus tested negative for every characteristic distinguished by Hektoen Enteric Agar.  Meanwhile, E. coli and P. fluorescens both seemed to turn out correctly, with development of orange and dark green hues (respectively).  These results implied that E. coli underwent lactose fermentation and P. fluorescens did not.
Triple Sugar Iron agar slants were possibly the most complex form of media used in this lab; incubation for 18-24 hours detected the presence or absence of H2S production, sugar fermentation, and/or gas production. These slants were commonly used by microbiologists to identify enteric bacteria, and contained 1% lactose, 1% sucrose, and 0.1% glucose concentrations.  Originally red in color due to the phenol red indicator, the agar slants were both stabbed and streaked with the inoculating needle.  This method formed two reaction areas in the same tube, one with deoxygenated/anaerobic conditions (the butt) and one with an oxygenated/aerobic environment (the slant).  A partially yellow (acid) butt and red (alkaline) slant meant glucose had been fermented, but coloration due to acid was not widespread because oxidative decarboxylation increases the pH of the slant.  A yellow butt and slant suggested that more than just glucose was fermented, leading to acid production throughout the medium.  An entirely red TSI tube indicated that no sugars had been fermented.  This last result never showed signs of gas production (splitting or bubbles in the agar) or hydrogen sulfide production (one or more black areas).  However, either of these characteristics could have been seen in a TSI tube with some amount of yellow coloration, because both were considered possible side effects of carbohydrate fermentation.  The gas/H2S production in TSI tubes was a byproduct of the reduction of thiosulfate to sulfite.
The Triple Sugar Iron tests seemed to turn out correctly in this lab, despite the many possibilities for human error.  The slant culture of P. vulgaris developed an acid slant and alkaline butt, which implied that a limited amount of carbohydrate fermentation had occurred, resulting in one of two possible byproducts:  hydrogen sulfide production.  E. coli’s slant was entirely acidic and yellow in color, a strong indication of carbohydrate fermentation.  Acid was not the only sign of this process, however, large amounts of bubbles and cracks in the agar lead to the assumption that gas had been produced as a byproduct.  TSI cultures of B. subtilis and P. fluorescens produced the same result:  an entirely alkaline tube without presence of H2S or gas.
The urease test detects for urease activity, or an organism’s ability to utilize urea.     These bacteria (seen in lab as P. vulgaris and P. fluorescens) make an enzyme that attacks the carbon-nitrogen bonds in amide compounds like urea.  This reaction produced CO2, NH3, and H2O, changing the originally straw-colored urease slant to a hot pink (positive test).  This hot pink color came from the pH indicator phenol red’s response to a sudden rise in pH from excess ammonia production.  Negative tests are still straw-colored.  As mentioned before, P. vulgaris and P. fluorescens both tested positive for urea utilization, producing the amide C-N bond-attacking enzyme to effect a swift color change in the urease slant to a bright pink.
Simmons Citrate Agar (SCA) tests determined a bacterial species’ ability to use sodium citrate as its only carbon source to produce energy.  If the species was equipped with a citrate permease to transport citrate into its cells, then the bacteria could convert the citrate to carbon dioxide and pyruvic acid.  This carbon dioxide combined with Na+ from the sodium citrate to produce alkaline sodium carbonate.  SCA tests also included NH4+ as a nitrogen source and bromothymol blue as a pH indicator.  Sodium carbonate production raises the pH and turns bromothymol blue from green to blue, indicating alkalinity.  Since citrate utilization requires oxygen, SCA tests are made in slant form.  Citrate negative tests normally should not have had presence of bacterial growth; however, this seemed to occur on both negative SCA results found in this lab (for E. coli and B. subtilis).  This could have been due to contamination from outside sources during inoculation, or a non-sterile SCA test tube.  P. vulgaris and P. fluorescens, on the other hand, both tested positive for citrate utilization, meaning they both were equipped with a citrate permease and could use sodium citrate as the sole carbon source.
Sulfide-Indole Motility (SIM) tests determined a bacterial colony’s amount of movement, hydrogen sulfide production, and indole production.  Indole was one of the byproducts of tryptophan hydrolysis, along with pyruvic acid and ammonia.  The NH4 and pyruvic acid got used for nutritional purposes by the bacteria, leaving an excess of indole (indicated by a formation of red coloration several minutes after the addition of Kovac’s reagent).

These SIM agar deeps were stabbed with the inoculating needle and incubated overnight, resultant appearances of tubes were examined to produce the following conclusions:  E. coli and B. subtilis both seemed to be motile bacterial species due to their apparent growth away from the line of stab inoculation.  Of the two, E. coli was the only one to reduce the thiosulfate in the tube to H2S, as was seen by the presence of black material in the tube.  P. vulgaris also produced hydrogen sulfide, but did not test positive for motility because the culture’s growth was restricted to the stab line.  According to the literature, however, this was incorrect, as P. vulgaris was a motile species.  The actual observation of PV’s test tube was probably mistaken, as the growth may not have been very far past the line of inoculation but was still enough to count as motile.  P. vulgaris’ indole test (performed by addition of Kovac’s reagent) also came out wrong, a false negative.  The likely cause of this error was that the Kovac’s reagent was not given enough time to react before observations were recorded two minutes later.  E. coli also had a false negative indole test result; meanwhile, the negative outcomes for P. fluorescens and B. subtilis were correct.  Observations of the SIM tubes after incubation lead to further conclusions that B. subtilis was motile but did not produce H2S, while P. fluorescens was neither motile or a thiosulfate reducer.

Methyl red tests were named after the pH indicator they contained, which developed a red hue if acidic end products of glucose catabolism were present.  Mixed acid fermenters acidified the medium by producing a variety of fermentation acids, which caused a greater fall in pH level than the effects of butanediol fermenters (production of butanediol, various organic acids, and acetoin).  Methyl red turns red (positive result) at a pH of 4, and yellow (negative result) at a pH of 6.  P. vulgaris and E. coli both had positive MR test results, with a switch to a red tinge after the addition of 4-5 drops of methyl red.  P. fluorescens and B. subtilis did not change in color once methyl red was added, but remained opaque (negative results).  These outcomes implied that P. vulgaris and E. coli both created acidic end products in their catabolism of glucose, while P. fluorescens and B. subtilis either did not ferment glucose or did not produce the same amounts of acidic products.

VI. References
(Harley JP, 2011). Laboratory Exercises in Microbiology 8th Edition: 139-143, 155-166, 195-197.

Lab 5: Bergey’s Manual

 

I. Purpose

The objective of this lab was to practice using Bergey’s Manual of Systematic Bacteriology to identify an unknown bacterial strain based on its biochemical properties alone.

II. Materials

The materials needed for this lab were as follows:
Bergey’s Manual of Systematic Bacteriology
- question sheet with detailed descriptions of two unknown bacteria

III. Methods

Identification of Unknown Bacteria

Bergey’s Manual of Systematic Bacteriology was skimmed through until a genus with similar or the same biochemical characteristics as one of the unknown bacteria on the question sheet.  The following sections of each article were used to identify the bacteria correctly:  name of genus, capsule description, enrichment, isolation, taxonomic comment(s), differential list of species within the genus, differences between the genus and other genera, and any further descriptive information.  These portions of each section were used to narrow down the possible identities of the unknown bacterial species from infinite to a definite match.  These definite matches (genus species names) were recorded as the answers to the questions on the question sheet.

IV. Results
The question sheet was turned in at the end of the lab, so there were no results to show here.
V. Discussion

Bergey’s Manual of Systematic Bacteriology was a fairly efficient method to identify bacterial unknowns from a comprehensive list of all biochemical characteristics that could be determined using standard differential tests.  There were four volumes in total, but only the ones pertaining to the bacterial species on the question sheet were skimmed through.  Introductory articles on bacterial taxonomy were located at the beginning of each volume for greater speed of identification.  The section of an article mentioned in the Methods section, “further descriptive information,” includes informative reads on topics pertaining to the genus, such as:  physiology, growth conditions, nutrition requirements, genetics, ability to cause disease, metabolism, ecology, and morphology.  These articles on bacterial genera include three main types of tables:  tables that differentiated between the species in the genus, tables that compared the respective genus to related genera, and tables that provided additional information on a particular species in the genus (possibly, this species is more commonly used in microbiology laboratories than others).

Lab 6: Biochemical Activities of Bacteria II, Unknown Identification, and Rapid Multitests
I. Purpose

The objective of this lab was to practice using rapid multitests such as the Enterotube II and API 20E to identify unknown bacterial specimens.  Also, the biochemical activities of various combinations of Bacillus subtilis, Escherichia coli, Enterobacter aerogenes, Pseudomonas fluorescens, Serratia marcescens, Lactobacillus casei, and Proteus mirabilis were tested through a series of tests, including but not limited to:  β-galactosidase metabolism, nitrate reduction, starch/casein/starch/gel hydrolysis, catalase, oxidase, litmus/lactose metabolism, glucose fermentation, H2S/gas production, and indole formation.

II. Materials
The materials needed for this lab were as follows:

 

– platinum inoculating loop
– normal inoculating loop
– inoculating needle
– zinc dust
– test tube rack
– Gram’s iodine
– API 20E Quick Index Booklet
– 1 5 ml Pasteur pipette with pipettor
– sterile mineral oil
– disposable gloves
– Barritts A reagent
– Barritts B reagent
– permanent marker
– Bunsen burner
– 10% ferric chloride
– biohazard container
– oxidase test reagent (catalase)
– distilled water
– 5 ml sterile 0.85% saline (capped test tube)
– API 20E system

~ strip

~ incubator tray

~ cover

– 1 starch agar plate
– 1 plate count agar with milk plate
– 4 cotton swabs
– 3 tryptic soy agar (TSA) slants
– 4 nitrate broth tubes
– Kovac’s reagent
– 1 BBL Enterotube II System
– Bunsen burner lighter
– 9 phenol red broth tubes with Durham tubes
~ 3 sucrose peptone broths
~ 3 lactose peptone broths
~ 3 dextrose peptone broths
– 4 litmus milk tubes
– 3 gelatin deeps
– Nitrate test reagent A
– Nitrate test reagent B
– Becton Dickinson Enterotube II Interpretation Guide
– 1.5% hydrogen peroxide
– 1% crystal violet solution
– 20% KOH (with 3% creatine solution)
– 5% α-napthol in absolute ethanol
– 1 tryptic soy agar culture plate of unknown bacteria
– tryptic soy agar cultures of:

~ Serratia marcescens

~ Enterobacter aerogenes

~ Bacillus subtilis

~ Pseudomonas fluorescens

~ Lactobacillus casei

~ Proteus mirabilis

 

III. Methods

Oxidase Test: Part I

    The cotton swab was aseptically inserted (clam-shelling) into the tryptic soy agar plate of unknown bacteria and dragged across the surface of the culture to pick up some bacterial cells on the tip.  Two drops of oxidase test reagent (catalase) were applied to the swab, and observations were made 20 seconds later.  If the bacteria on the swab turned blue by 20 seconds after the application, the bacteria tested positive for cytochrome oxidase production.  If the bacteria did not change color, it was considered an oxidase negative test result.  If oxidase test results came out negative, this unknown bacterial culture was then placed to the side for later use in inoculation of the Enterotube II and API 20E systems.  If not, a different unknown bacterial culture had to be found and tested.

Inoculations

    Aseptic technique (relating to plates, inoculation instruments, and tubes) was strictly followed in all bacterial transfers performed in this lab, to avoid contamination.  Starch agar plates were inoculated with a single straight streak about 1” in length for each bacterium (BS, EC, and PM), using an inoculating loop.  The loop was also used for inoculating the casein agar plates; however, instead of a line, a circle-shaped spot inoculation about .5” in diameter was performed in each sector of the plate using EA, EC, and PF.

Tubes with liquid medium (including litmus milk, phenol red, and nitrate) were inoculated by swirling a loopful of the respective bacteria around in the broth.  Litmus milk tubes were inoculated with LC, EC, SM, and PF.  Nitrate broths were inoculated using SM, EC, PF, and PM.  Phenol red tubes were inoculated by categories, with each bacteria being used once for sucrose, dextrose, and lactose tubes; PM, EC, and EA were utilized for these tests.

Tryptic soy agar slants were inoculated with a single zigzag streak of an inoculating loop across the surface of the slant.  Gelatin deeps were inoculated with a single stab from an inoculating needle.

The Enterotube II was inoculated by unscrewing the caps at both ends, dragging the needle through an unknown oxidase negative bacteria culture, and using the handle to simultaneously twist and pull the needle all the way through every compartment, without completely removing the needle.  The needle was then pushed back into the tube until the tip of the needle was in the H2S/indole compartment, at which point the rest of the needle was broken off and used to poke ventilation holes in the plastic covering of the ADON, LAC, ARAB, SORB, VP, DUL-PA,UREA, and CIT compartments.

The API 20E system was inoculated according to the “First Period” procedure in Exercise 35 of the lab manual.  All inoculated media were loaded into/onto a test tube rack and incubated at 35oC for 18 – 24 hours.

———————————————— COMEBACKS ————————————————-

 

Enterotube II

    The results for the Enterotube II system were interpreted with regard to Figure 36.1 from the lab manual.  The indole test was performed by addition of 2 drops of Kovac’s reagent to the H2S compartment, using one of the Pasteur pipettes.  Results were determined after 10 seconds.  The Voges-Proskauer test was performed with the addition of two drops of 20% KOH and three drops of α-naphthol.  Results for the VP test were determined after 20 minutes.  All outcomes were recorded on the data sheet for later analysis.

API 20E

    The results for the API 20E system were interpreted according to Exercise 35 in the lab manual.  If the GLU microtube resulted in a negative glucose test, then the unknown was not a member of the Enterobacteriaceae and testing could not be continued.  A positive test (yellow, presence/absence of bubbles does not matter) meant that the following reagents had to be added in the order listed.  One drop of 10% ferric chloride was added to the TDA microtube (positive test = red-brown color, negative test = yellow color).  One drop of Barritts A and one drop of Barritts B were added to the Voges-Proskauer microtube.  Barritts B was added first, and results were observed after 10 minutes (positive = red-pink color change, negative = no color change).  One drop of Kovac’s reagent was added to the IND microtube; a positive indole result was shown by formation of a red ring within two minutes of addition of the Kovac’s.  A negative result was indicated by the formation of a yellow ring after the allotted time had passed.  The GLU tube was then examined for gas bubbles, with a positive result being the presence of bubbles and a negative result, the absence.  Two drops of nitrate test reagent A and two drops of nitrate test reagent B were added to the GLU microtube.  A positive test result showed after 2-3 minutes as the development of a red hue; a negative test involved the forming of a yellow hue after 2-3 minutes.  If the test turned out negative, a speck of zinc dust was added to the tube.  Ten minutes later the tube was observed for the development of a orange-pink color (negative nitrate reduction test result).  If the color changed to yellow instead, nitrogen reduction had occurred and nitrogen gas had been produced.  Finally, one drop of hydrogen peroxide was added to the MAN, INO, and SOR cupules.  A positive catalase test result involved the appearing of gas bubbles within two minutes.  A negative test has no such bubbles.  All outcomes were recorded on the data sheet for later analysis.

Starch Hydrolysis

    Several drops of Gram’s iodine were added to the bacterial streaks on the starch agar plate.  Any bacteria that seemed to repel the iodine, creating a clear area around the streak, were considered to have gone through starch hydrolysis (positive outcome).  Bacterial streaks that did not form a clear region around the line of growth, or beneath which the medium turned blue with application of iodine, did not hydrolyze starch (negative result).  All outcomes were recorded on the data sheet for later analysis.

Gelatin Hydrolysis

    After incubation, the tubes were tilted at a slight angle for observation of gel consistency.  If the nutrient gelatin was liquefied, gelatin hydrolysis had occurred and the bacteria in that culture was marked down as a positive result.  Solidified gelatin meant that no hydrolysis had occurred; this was noted as a negative outcome for that bacterial species.  At least the uninoculated control, if nothing else, should have tested negative.  All outcomes were recorded on the data sheet for later analysis.

Casein Hydrolysis

    Casein agar plates were examined for presence or absence of clear zones around the bacteria, called “zones of proteolysis.”  Presence of a zone indicated a positive casein hydrolysis test; the bacterial species in question was able to liberate proteases.  Absence of this clear zone meant a negative result, and the species was not able to hydrolyze casein or liberate proteases.  All outcomes were recorded on the data sheet for later analysis.

Oxidase Test: Part II

    Cotton swabs were used to pick up a small amount of each bacteria (one species per swab) growing in the TSA tubes that would later be used for catalase tests.  One to two drops of oxidase reagent were applied to each swab, and results were observed after about 20 seconds.  Like the first oxidase tests run in this lab, blue color change meant a positive result and no color change meant a negative result.  All outcomes were recorded on the data sheet for later analysis.

Catalase Slants

    Several drops of 3% hydrogen peroxide were added to each tryptic soy agar slant.  If gas bubbles appeared, the test was positive.  If no bubbles formed, results were found to be negative.  All outcomes were recorded on the data sheet for later analysis.

Litmus Milk Tubes

    After incubation, the four litmus milk tube cultures were observed for color change and curd formation at the base of the tube.  Gas production as a side effect of fermentation was also possible, but clear results (cracks or bubbles in the medium) were difficult to see.  If litmus had been fermented, the milk turned pink (positive test).  A blue-purple coloration indicated that lactose fermentation had not taken place (negative fermentation result), but protein metabolism had.  (the blue color comes from the proteins’ alkaline pH)The tube may also be white in color, from complete reduction of litmus contents.  Curd at the bottom of the tube signified the presence of casein digestion.  All outcomes seen in lab were recorded on the data sheet for later analysis.

Nitrate Broth Tubes

    Presence of gas bubbles in the nitrate broth tubes meant that the bacteria in the respective tubes had reduced the nitrate in the broth to nitrogen gas.  Absence of these bubbles could still have meant nitrate reduction had taken place, but to a nongaseous end product such as nitrite.  Five to ten drops of nitrate test reagent A and five to ten drops of nitrate test reagent B were added to each tube and mixed by finger-vortexing.  Results were almost immediately observable; a pink-red color was a positive nitrate test result and an absence of color meant a negative result.  Negative tests were confirmed by adding several grains of zinc powder or 5-10 drops of nitrate reagent C and shaking the tube.  Results from negative test confirmations were observed after 8-10 minutes.  If nitrate was present in the medium, then the broth appeared red at this point.  All outcomes were recorded on the data sheet for later analysis.

Phenol Red Tubes

    Red coloration of tubes after the incubation period could either mean no fermentation took place (no gas bubble in Durham tube) or alcohol fermentation occurred (gas bubble in Durham tube).  Any amount of yellow hue implied acid production, and any size bubble in the Durham tube meant that gas had been produced.  Respective positive and negative results were recorded on the data sheet for later analysis.

IV. Results

Phenol Red Tubes

sucrose

dextrose

lactose

gas

acid

growth

gas

acid

growth

gas

acid

growth

P.  mirabilis

+

+

+

+

+

+

E. coli

+

+

+

+

+

+

+

+

+

E. aerogenes

+

+

+

+

+

+

+

+

+

Gelatin Deeps

growth

gelatin hydrolysis

P. mirabilis

+

+

E. coli

+

B. subtilis

+

+

CONTROL

 

Catalase Slants & Oxidase Test

growth

gas

oxidase

E. aerogenes

+

+

E. coli

+

+

P. fluorescens

+

+

+

 

Enterotube II

glucose

lysine

ornithine

H2S

indole

adonitol

lactose

arabinose

sorbitol

VP

dulcitol

PA

urea

citrate

Unknown

+

+

+

+

+

+

+

+

+

+

+

+

 

 

 

 

 

Nitrate Broths

growth

nitrate reduction

S. marcescens

+

+

E. coli

+

+

P. fluorescens

+

+

P. mirabilis

+

+

 

Starch Agar Plate

growth

starch hydrolysis

B. subtilis

+

+

E. coli

+

P. mirabilis

+

Litmus Milk Tubes

growth

fermentation

curd

L. casei

+

+

+

E. coli

+

+

+

S. marcescens

+

+

P. fluorescens

+

+ (tiny spot)

 

Casein Agar Plate

growth

casein hydrolysis

E. aerogenes

+

E. coli

+

P. fluorescens

+

+

 

 

 

 

 

 

 

 

V. Discussion
This microbiology lab involved the use of rapid multitests (the Enterotube II and API 20E systems) to identify unknown bacterial specimens.  The biochemical activities of Bacillus subtilis (BS), Escherichia coli (EC), Enterobacter aerogenes (EA), Pseudomonas fluorescens (PF), Serratia marcescens (SM), Lactobacillus casei (LC), and Proteus mirabilis (PM) were studied via inoculation of various differential and/or selective cultures.

The API 20E System used in this lab was a fast, standardized, more managable test that combined conventional biochemical procedures used to identify 127 various taxa of bacteria in the Enterobacteriaceae family and other gram negative groups.  The system used microtubes to perform 22 common biochemical tests on pure cultures, which were standard to most identification procedures.  The structure of the system itself consisted of 20 chambers, each made of a microtube and depression known as a “cupule.”  Cupules helped to create anaerobic conditions – which were required for certain tests – via the addition of mineral oil, to block off the oxygen supply.  The API 20E strip is read by recording color changes after incubation; for some indicator systems, reagents were added before determining outcome.  Unknown bacteria were identified by using a chart technique similar to that on page 225 of the lab manual (questions 1 and 2) to give the unknown a seven digit profile number.  The API 20E Profile Index Booklet could then be consulted to find the bacterial name that matches up with the profile number.

Once results were completely formed for the API 20E strip and recorded, the profile was calculated to be 0774770.  The first three digits were the most crucial to the identification of the unknown, calculating seven was not entirely necessary.  The unknown culture was classified as Proteus mirabilis using the API 20E Profile Index Booklet.

The Enterotube II System was very similar to the API 20E in that it theoretically accomplished the same results.  However, inoculation of the Enterotube went much more quickly than that of the API 20E System.  The Enterotube was not as inclusive as the API 20E, though.  Only glucose-fermenting, gram negative, oxidase negative members of the Enterobacteriaceae family could be identified using the Enterotube.  The tube was divided into 12 compartments, each with a different solid agar culture medium.  Some were aerobic, and had air holes to allow the flow of oxygen to reach bacterial cells, but most were conducted in anaerobic conditions.  Enclosed in the tube was an inoculating needle specially made for the Enterotube, allowing for speedy inoculation of all 12 compartments in two swift twisting motions.  After 18-24 hours of incubation, the color changes in each compartment were recorded and interpreted according to Table 36.1 in the lab manual.  Some compartments had reagents added to them to produce the final test result, and this is what was recorded in the Results section for those particular compartments.  These results were placed into the chart from question 1 of Exercise 36 in the lab manual to obtain a five digit number that can be compared to the Enterobacter II Interpretation Guide to identify the unknown bacteria.

The Enterotube II System performed on the unknown bacterial culture in this lab was successfully carried out; results for each compartment were recorded in the Results section.  The code was calculated from the results to equal 37347, which did not match any numbers in the Enterobacter II Interpretation Manual.  Therefore, the true identity of the unknown bacteria defaulted back to the result from the API 20 E multitest system.

Phenol red broths were used to determine one of the most commonly studied biochemical characteristics of bacteria:  the ability to ferment carbohydrates.  There were three different carbohydrates used as indicators of beta-galactosidase activity in bacterial cultures.  These were known as sucrose, dextrose, and lactose; three phenol red tubes were made with each to accommodate for the three different types of bacteria used to inoculate the phenol red broths (PM, EC, and EA).  Any amount of yellow coloration (from phenol red reacting to acidic pH) in the resulting tubes after 24-48 hours of incubation indicated the presence of fermentation products.  Gas production could also be identified by the phenol red tubes, because any amount produced would be trapped inside the Durham tube.  As noted in the methods, complete red coloration of a gas-less tube denoted complete lack of fermentation, as the phenol red had not indicated any lowering of pH by acidic products. PM fermented carbohydrates in all but the lactose tube, but EC and EA fermented in all phenol red tubes.  At first glance, this could possibly have been explained by a false negative result (from an excessively large inoculum).  However, further research revealed that PM cannot ferment lactose, so the negative result was proven correct.

Gelatin hydrolysis was seen as a rather important biochemical characteristic because it could be used to assess pathogenicity of some bacteria.  Those bacteria that could hydrolyze gelatin were more likely to cause disease than those who could not, because the production of gelatinase could be correlated with the ability of a bacterium to break down tissue collagen and become more virulent.  PM and BS both tested positive for gelatin hydrolysis, while EC returned a negative test.  Of course, this result could have been an exception to the “usually more pathogenic” rule, because EC has long been known to be more pathogenic than both BS and PM.

Nitrate broths determined the ability of a bacterium to reduce nitrate.  Nitrate could be reduced into gaseous nitrogen, forming a bubble in the Durham tube, or reduction could have stopped at nitrite and other nongaseous products.  Therefore a confirmation of negative results (after addition of nitrate reagents A and B) was needed to be sure that nitrogen fixation was definitely not occurring.  This was done through the addition of nitrate reagent C and observing results after 5-10 minutes.  SM, PF, and PM all produced immediate positive (red) results for occurrence of nitrate reduction, yet EC remained clear (negative result).  A recheck was done to make sure that no false negative had occurred, and EC also tested positive after a period of eight minutes.  Therefore all nitrate broths brought back positive results for nitrate reduction, and EC, PM, PF, and SM were all confirmed to use nitrate as a terminal electron acceptor during anaerobic respiration.

The test for catalase presence in EA, EC, and PF was performed on tryptic soy agar because of its general properties as a nutrient and growth medium.  According to the results, neither EC, PF, or EA were strict anaerobes (rather they were obligate aerobes or facultative anaerobes), which corresponds with previous scientific findings.  Catalase production/activity was tested for by the addition of hydrogen peroxide to the slants, formation of bubbles meant a positive test.  Catalase catalyzed the destruction of hydrogen peroxide, which was harmful to the respiration classes that these bacterial species fell under.  When the oxidase test was performed, however, only PF produced positive results (presence of oxidase turned swabful of bacteria blue).  According to the lab manual, these oxidase enzymes in PF play a key role in the function of the electron transport system when PF cells respire aerobically:  “Cytochrome c oxidase uses oxygen as an electron acceptor during the oxidation of reduced cytochrome c to form water and oxidized cytochrome c.”

As for the casein agar plates, these differential media detected zones of proteolysis, or casein hydrolysis, around the species of bacterial colonies that were able to secrete proteolytic enzymes.  This plate was one of the most easily conducted biochemical characteristic-determining tests in this experiment, because it did not require addition of reagent or specific timing for result determination to be viable.  EA, EC, and PF all grew successfully on the agar, but only PF formed a zone of proteolysis.  Therefore PF was the only bacteria to be tested for casein hydrolysis in this lab that came back with positive results, indicating PF went through a hydrolytic reaction that produces soluble amino acids, which were then transported into the cell through proteases and catabolized to produce energy.

The starch agar plates were also a relatively simple test; all that was required was the addition of a small volume of gram’s iodine to yield immediately visible results.  Both EC and PM failed to hydrolyze starch and repel the Gram’s iodine, but BS was successful on this front.  As a result, BS was found able to rapidly hydrolyze the two constituents of starch:  amylose and amylopectin.  This hydrolysis yielded dextrins, glucose, alpha-amylase, and maltose.  The alpha-amylase is what repels the iodine and produces the clear region surrounding the starch hydrolysis-capable bacterial colony.

Litmus was a pH indicator used in this lab that was blue-purple under alkaline conditions and light pink in acidic conditions.  This pH indicator is the reason for the title of “Litmus Milk” tubes.  The milk part of the name comes from the tube’s milk-like consistency and the fact that it contains casein and lactose.  LC showed up for the first, last, and only time in the experiment in one of the litmus milk tubes.  EC, PM, and PF were used for inoculating the other three tubes.  LC, EC, and PF all produced a solid “curd” product and the base of their respective tubes, indicating casein metabolism.  However, PF did not obtain positive fermentation results, and remained blue in color.  LC, EC, and SM all went through lactose fermentation and caused litmus to change the tubes to a pink hue.  Peptonization – in which the casein would be metabolized completely into amino acids – did not occur with any of the bacteria in this experiment, but if it had, a clear brown liquid would have resulted.
VI. References
(Harley JP, 2011). Laboratory Exercises in Microbiology 8th Edition: 132-136, 147-150, 169-179, 189-192, 213-229.

pGLO Lab Analysis

pGLO Lab
By: Yi Yu

Abstract

            Plasmids are pieces of circular DNA that are in bacteria which can code for genes that can cause the bacteria to have extra “features”. With the pGLO plasmid this extra “feature” causes bacteria to glow (from the Green Fluorescent Protein that was inserted) and also has resistance to ampicillin (from the beta-lactamase protein). By adding the pGLO plasmid the E.coli bacteria will be able to grow on agar with arabinose sugar and ampicillin. The main reason for doing this lab is to better understand how genes control traits, they best way to do this is by doing mutagenesis. There are a few different ways to preform mutagenesis and in this lab it is done by adding a transposon to the pGLO plasmid and then looking at how the transposon effects how the bacterium looks and grows. By looking at how the bacteria grows and looks we can determine where the transposon inserted; this helps us better understand how genes control traits. In the sample we worked with the mutagenized bacteria did not glow green under UV light which tells us that the transposon inserted into the araC gene, and is confirmed in the electrophoresis gel.

Introduction

            Plasmids have been studied in genetic research since they were discovered in 1952 by Joshua Lederberg. They have been used to figure out how certain bacteria reacts to different types of antibiotics and to change how bacteria functions by adding things into the plasmids. In this lab the plasmid that we focused on was the pGLO plasmid. This plasmid has three main coding regions that we will be looking at. The first gene region is the region that encodes for the beta-lactamase (bla) enzyme. “The beta-lactamase protein is produced and secreted by bacteria that contain the plasmid. Beta-lactamase inactivates the ampicillin present in the LB nutrient agar, allowing bacterial growth. Only transformed bacteria that contain the plasmid and express beta-lactamase can survive on plates that contain ampicillin.” (Leatherman 2011). The Green Fluoresce Protein (GFP) is a gene code that is inserted into the pGLO plasmid to make it glow green under UV light with the help of the araC gene. The araC gene encodes a protein that binds to arabinose and the promoter region as well in the presence of arabinose, when bound to the promoter region of the GFP gene it translates the protein that will make the bacteria glow green.

Mutagenesis does not happen very often in nature but can be used regularly in genetics labs to help study different functions of bacteria and plasmids. By doing mutagenesis on bacteria we can better understand how bacteria translate genes and how they react to their environment when something is changed. There are a couple ways to preform mutagenesis like using chemicals or X-rays. In this experiment mutagenesis is performed by inserting a transposon into the pGLO plasmid. The Tn5 transposon will be inserted into the pGLO gene; the Tn5 transposon encodes for a protein that leads to resistance of kanamycin, which will help us see if the transposon inserted correctly.

After the transposon is inserted into the pGLO plasmid, it will be taken up by E.coli bacteria through transformation. Transformation is when bacteria take in free DNA from their surroundings. Once the plasmids were taken in the bacteria was grown on nutrient agars with different combinations of ampicillin, kanamycin, and arabinose along with control groups to see how the mutant bacteria was affected by the transposon.

Bacteria use restriction enzymes on their own to get rid of bacteriophages by either cutting the proteins that they make or by cutting the phage directly (Aude). We are able to use these restriction enzymes to do restriction enzyme digest to cut the plasmids that were inserted into the bacteria. “Restriction enzymes are commonly used to cut DNA at particular nucleotide sequences. Often this is to make a new “recombinant piece of DNA, and is followed by ligating two pieces together.” (Leatherman 2011). The plasmids were cut at known locations of the plasmid with Nde 1 and EcoR1 enzymes. Knowing the amount of base pairs that are in the plasmid (5,400 bp) and where the restriction enzymes cut we can examine the different pieces of the cut plasmid to find out where the transposon inserted.

pGLO plasmid restrictino enzymes

Methods

            Use the Tn5 insertion kit to insert the Tn5 transposon into the pGLO plasmid. The first step was to mix 6.4 microliters of water, 1 microliter of EZ Tn5 10X reaction buffer, 1 microliter of .2 micrograms/microliter of pGLO plasmid, 0.6 microliters of Tn5 transposon, and 1 microliter of EZ-Tn5 transposase (helps transposon insert) in this order and mix by vortexing, then centrifuge. Incubate tube at 37 degrees Celsius for 50 minutes, then take tube out and spin again and return tube to incubator for another 50 minutes. Spin once more and add 1 microliter of stop solution to tube and mix well and spin liquid to bottom of the tube again. Heat tube to 70 degrees for 10 minutes; store in freezer for next time.

Transfer 1.5 mL of E.coli to three microcentrifudge tubes and put on ice for 10 minutes. Centrifuge tubes for 1 minute and get rid of all supernatant. Add 300 microliters of ice-cold CaCl2 to all three tubes; dissolve the pellet completely by pipetting and place tubes on ice for 2 minutes. Centrifuge tubes for 1 minute and discard the supernatant. Add 60 microliters of CaCL2 to each tube again and gently re-suspend the pellet and put on ice for 10 minutes. Label the three tubes, No DNA, pGLO wt, and pGLO mutant. Add nothing, 1 microliter of pGLO DNA, and 1 microliter of Tn5 mutagenesis pGLO reaction to each tube respectively. Gently mix tubes by tapping them and place on ice for 15 minutes. Heat shock the tubes at 42 degrees for exactly 90 seconds and return to ice immediately for 2 minutes. Add 1 mL of LB broth to tubes then transfer contents of each tube to a test tube and incubate at 37 degrees for 30 minutes with shaking. Obtain and label 5 different agar plates like shown in table 1 and label them. Transfer bacteria into new microcentrifudge tubes and spin them for 1 minute and remove 500 microliters of supernatant, re-suspend bacteria in broth. Add 200 microliters of each reaction to their respective agar plates and spread bacteria over the agar plates. Incubate plates at 37 degrees in prep room for a day.

DNA

Type of plate

Prediction

No DNA Nothing added Lawn of white
No DNA Ampicillin No growth
pGLO wild-type Ampicillin White colonies
pGLO wild-type Ampicillin + Arabinose Green colonies
pGLO mutagenized Kanamycin + Arabinose Green + White colonies

Table 1. DNA added to the different plates and predictions.

Prepare two sterile culture tubes with 5 mL of LB broth, one with ampicillin and one with kanamycin, label which is which. Examine plates and choose on wild-type colony and put colony into the LB broth with ampicillin. Look at plate with the pGLO mutant and look to see if any of the colonies glow green under UV light. Colony that was picked for this experiment was white. Take colony and put in LB broth with kanamycin. Put tubes into incubator at 37 degrees and incubate overnight. Look at all plates and compare the results with your predictions. Get cultures from the incubator and transfer them into 15 mL tube to centrifuge. Spin in refrigerated centrifuge at 3000 rpm for 5 minutes and discard supernatant. Add 250 microliters of P1 buffer (removes RNA) to each bacteria pellet and re-suspend the pellet, and immediately transfer to microcentrifudge tube. Add 250 microliters of P2 buffer (lysis of cells) and mix by inverting for no more than 5 minutes. Add 350 microliters of N3 buffer (pH ideal for column binding) and mix immediately by inverting the tube. Spin for 10 minutes until there is a white pellet that forms. Pipet supernatant into separate spin columns and spin for 30 seconds then discard flow-through. Add 750 microliters of PE buffer (washes DNA) to column and spin for 30 seconds and discard flow-through. Place column in new 1.5mL microcentrifudge tube; add 50 microliters of water (elution of DNA) to spin column and let it stand for 1 minute. Spin for 1 minute then discard spin column and keep the flow-through. Measure the concentrations of the two samples of DNA, then put in freezer for next week. Concentration of wild-type DNA was 68.2 nano grams/microliter with a purity of 1.84 and the mutant DNA had a concentration of 81.9 nano grams/microliter with a purity of 1.64 which was not as pure

            Use the restriction map to determine the number of bands in the wild-type plasmids cut at the different places.

Uncut Nde1 EcoR1 Nde1 + EcoR1
Number of bands and their sizes 5,400 bp 3,300 bp1,800 bp

300 bp

5,400 bpNot circular 2,800 bp1,800 bp

500 bp

300 bp

Table 2. Band sizes of wild-type plasmids with different cuts.

Mix each solution based on what each tube needs. Use the concentration of DNA to determine how much DNA needs to be added to each tube (500/concentration=how much DNA is need).

Uncut wt Uncut mutant Nde1 wt Nde1 mutant EcoR1 wt EcoR1 mutant Nde1 + EcoR1 wt Nde1 + EcoR1 mutant
Water 12.67 13.9 10.17 11.4 10.17 11.4 10.12 11.4
10X buffer None None 2Buffer 4 2Buffer 4 2 EcoR1 buffer 2 EcoR1 buffer 2 EcoR1 buffer 2 EcoR1 buffer
DNA: 5 milligrams 7.33 6.1 7.33 6.1 7.33 6.1 7.33 6.1
Nde1 enzyme (20units/microliter) None None .5 .5 None None .5 .5
EcoR1 enzyme (20units/microliter) None None None None .5 .5 .5 .5

Table 3. Mixtures of each tube. (all in microliters)

Place the eight tubes in incubator at 37 degrees for 1.5 hours for reaction to proceed, once 1.5 hours is up put tubes in freezer for next time.

Dilute 50X buffer to 1X buffer; add 20 mili liters of 1X TAE buffer in a graduated cylinder and then add 980 mili liters of water to the cylinder. Make a 1.2% agarose gel; weigh out .6 grams of powder agarose and empty into a flask, then take 50 mL of 50X TAE buffer and add it to the flask. Mix the agarose and the buffer then put it into a microwave for about a minute or until the solution becomes clear. Let the flask cool until it you can touch the flask without burning yourself.  Set the tray in the running box with both side up and the comb in place and pour the agarose solution into the tray. Let the agarose sit until it becomes a milky white color and hardens. While waiting load 2 microliters of 10X loading dye to each tube and mix by pipetting. Lower the sides of the tray and then fill the running box with the TAE buffer until it covers the top of the gel. Load the 1kb ladder on the far left end with 10 micro liters of 1kb ladder. Take the PCR reactions from the previous week and fill in the next 8 spaces in the gel with 20 micro liters of the restriction digest reactions in order. After the gel is loaded run the gel for about 45 minutes at 100 volts, check periodically to see if the dyes are moving down the gel. Once the dye has gone about half way down the gel take it out of the running box and place the gel in a staining box. Pour SYBR green into the staining box until the SYBR green covers the top of the gel, swirl the box every 5 minutes. The SYBR green binds to the DNA and allows us to look at the DNA under a UV light. After 15 minutes take the gel out of the box and pour the SYBR green back into its bottle. Take your gel and place it in a UV box and take a picture of your gel to see the results of your PCR.

Results

            When the bacteria was first examined the mutagenized bacteria did not glow green when put under UV light which means that the transposon could have inserted itself into either the araC gene or into the GFP gene.

DNA

Type of plate

Results

No DNA Nothing added Lawn of white
No DNA Ampicillin 13 colonies
pGLO wild-type Ampicillin White colonies over 100
pGLO wild-type Ampicillin + Arabinose Green colonies over 100
pGLO mutagenized Kanamycin + Arabinose 8 all white colonies

Table 4. Results of the agar plates.

            When mixing the restriction enzymes, the restriction enzyme was mistaken for buffer when being added to the mutant tubes which lead to the mutant tubes to be unreadable on the gel.

pGLO Lab

Figure 2. Row 1 is the 1kb ladder. 2 wild-type uncut plasmid. 3 wild-type nde cut. 4 wild-type eco cut. 5 wild-type eco/nde cut. 6 mutant uncut. 7 mutant nde cut. 8 mutant eco cut. 9 mutant eco/nde cut

From this gel we can see that the wild-type uncut and EcoR1 cut are the same which is what was expected. There are four bands on the wild-type EcoR1/Nde1 cut in the 300 and 500 regions and in the 1000 and 2000 areas. And three bands on the Nde1 cut in the 300, 1000, and 2000 areas. So all of the wild-type samples were where they were predicted to be. Unfortunately the mutant samples are not readable.

plasmid gel 2

Figure 3. Row 1 is the 1kb ladder. 2 wild-type uncut plasmid. 3 wild-type nde cut. 4 wild-type eco cut. 5 wild-type eco/nde cut. 6 mutant uncut. 7 mutant nde cut. 8 mutant eco cut. 9 mutant eco/nde cut.

In this figure you can see that in row 6 it didn’t travel as far as the wild-type uncut in row 2 telling us that the transposon is in the sample. Looking at row 7 and 9 there is a missing bar at around 1,800 base-pairs like in rows 3 and 5. This is where the transposon is inserted itself into to the araC gene which is in one of the Nde1 cuts that it about 1,800 base-pairs. In the gel the addition of the transposon to this region causes the 1,800 bar to stay back to where the 2,800 base-pair bar is and makes it look like just one bar on the gel.

Discussion

            The transposon was found in the araC gene of the pGLO plasmid from the gel results. With the transposon being in the araC gene it doesn’t make the protein that binds to the promoter region of the GFP gene which resulted in the mutagenized bacteria to not glow green under the UV light. The insertion of the transposon in the araC gene might also explain why there was so little mutant growth on the plate the mutant bacteria was placed on. The problem with the original gel was that more enzyme was added to the tubes which made the mutant samples unreadable on the gel.

 

References

Aude A Bourniquel, Thomas A Bickle, Complex restriction enzymes: NTP-driven molecular motors, Biochimie, Volume 84, Issue 11, 1 November 2002, Pages 1047-1059, ISSN 0300-9084, 10.1016/S0300-9084(02)00020-2.

Leatherman J. 2011 Genetics Laboratory Manual. University of Northern Colorado

Effects of a Frequently Disturbed Industrial Area on Leaf Litter Invertebrate Communities

By: Mark Nie, Ricky Snow

Abstract

            Soil Invertebrates are an important part of the ecosystem in which they reside.  They are often overlooked because of the fact that they are so small and cannot be seen with the naked eye.  The following study looks at the species richness of soil invertebrates while comparing two different parks that were in completely different areas.  The parks that were compared are Pioneer Park in Kenton County, Kentucky and the second growth of Withrow Nature Preserve in Cincinnati, Ohio.  The hypothesis that we were testing was that the more disturbed, industrialized Pioneer Park would have a lower species richness compared to the less disturbed, more rural Withrow Nature Preserve.  The method for testing this was taking two different samples of leaf litter from within each park that measured 10cm by 10cm for a total of 40cm2.  These samples were then put in a Berlese funnel for a full 24hrs and collected in a dish of alcohol.  After going over the samples from each park, we found that the impact of an industrialized area had no effect on the species richness of the environment.  Further data collection would be needed to corroborate whether this would also be true over a larger area.

 

Introduction

            Leaf litter and soil invertebrates are very important to the overall ecosystem that they are a part of by playing a major role in nutrient cycling. Being near the base of the trophic web, these soil invertebrates can be important bioindicators. The species richness of these soil invertebrates could be used to determine the overall health of the surrounding ecosystems.

The question this study is aiming to answer is what effects a frequently disturbed industrial area has on nearby ecosystems by looking at the soil invertebrate communities’ species richness. In order to determine the health of the area in question samples should be collected at both the site in question and a control site, then the two communities can be compared. The hypothesis states that a frequently disturbed industrial area will have a low species richness compared to a less disturbed area without industrial influence.

 

Methods

            The first site we tested was Pioneer Park located in Kenton County, Kentucky. The park is located along a major highway near several factories and residents. The park has little forested area which is primarily new growth and is a heavily used recreational park that includes sporting fields, a dog park and picnicking areas. Banklick Creek runs through the park and is downstream of residential areas, the park is frequently flooded after heavy rainfalls.

The second site we tested was Withrow Nature Preserve located in a rural area west of Cincinnati, Ohio. The preserve has a history of farming and is heavily forested including old growth as well as new growth and is primarily used for its hiking trails. The preserve is much less disturbed than Pioneer Park, the preserve also lacks nearby industrial influence as well as a lack of a nearby creek.

The sampling technique used was very simple and rudimentary, requiring nothing more than a few meter sticks and grocery bags.  The samples that were collected were in two 10 cm2 sections for a total of 40 cm2 at each site. At each site the sections collected were in new growth forest one near the edge of the forest and the other deeper in the forest. The sample from Pioneer Park included species such as sycamore, oak and maple leafs among others and Withrow Nature Preserve sample consisted of primarily oak and maple leafs. Both sample sizes were similar as well as presence of woody material. The leaf litter within the measured sections ware picked up and placed in a plastic bag to be brought back to the lab and placed in a Berlese funnel for 24 hours where a light in the top of the funnel would drive the organisms within the leaf litter out the bottom of the funnel into a jar containing 70% ethanol. The organisms were then separated into general species and counted the results of which were then analyzed for species richness.

 

Results

            The results of the study were unexpected compared to what we were looking for.  As seen in Table One in Appendix A, Pioneer Park had a density of 44 organisms.  These organisms were largely dominated by one species of springtail accounting for almost 50% of the total organisms collected.  Withrow Nature Preserve had a few more organisms but overall, the density was close to that of Pioneer with 53 total organisms as depicted in Table Two also in Appendix A.  This sample was dominated sheer number wise by mites, which accounted for 30% of the total, but species diversity wise, was dominated by springtails; there were three different species within this sample alone.  In Appendix A, there are rank abundance graphs for each park, which show the sheer numbers for each organism we found within the study at each site.  Our calculated Shannon-Weiner Diversity Index for Pioneer Park was 4.526 and 4.527 for Withrow Nature Preserve.  Overall, each site was pretty similar when it came to species richness and diversity of organisms within the samples.

 

Discussion

            The results of our study in a way threw us for a loop because they were so unexpected from what we thought they were going to turn out to be.  When looking at the figures and tables that we put together, there was a very small density difference between Pioneer Park and Withrow Nature Preserve.  Pioneer Park had an overall density of 44 organisms and Withrow Nature Preserve had an overall density of 53 organisms.  Apart from a few minor differences in how the organisms looked, the samples had very similar species comparisons as well which was interesting.  One reason we were so confident about our hypothesis was because we were looking not only at two different parks in different areas but also in two different states.  However, it appears as if the location, at least for our small study, had little to no effect on the density of species richness when it came to the different sites.

The biggest result that came out of our study was the Shannon-Weiner Diversity Index that we calculated for each site.  The calculated index for Pioneer Park was 4.526 which one the scale is an indication of a very diverse and rich area.  For Withrow Nature Preserve, the index that was calculated was 4.527, which again is a very diverse and rich area.  However, what is interesting here is how close these numbers are.  The fact the indices are .001 away from each other disproves our entire hypothesis that there would be an impact based on the industrialized location of Pioneer Park compared to Withrow Nature Preserve.  As indicated by the indices, both locations are very diverse and species rich which is a positive aspect that came out of our study.

Our study had obvious limitations that would need to be corrected to further our study.  One limitation of our study was the size of the plots we collected from and the number we collected.  We only collected 40cm2 at each location and only acquired two samples for each as well.  For locations that are as big as Pioneer Park and Withrow Nature Preserve, we would need to not only widen the area we collected from but also collect a vast more number of samples.  Another limitation that was present was the fact that we only collected from two different parks that were local.  If we were to add more parks in different areas, we would be able to further test our hypothesis on these parks in comparison to what we found now.  The addressing of these limitations would help to further our study and possibly make our hypothesis more valid.

 

Acknowledgements

            We would like to thank one another for the help that was given in this study.  Each of us brought a different overall creative idea to make the study unique and successful.  Also, we would like to thank Dr. Shannon Galbraith-Kent, professor of biology at Thomas More College, for her help with the study.  Dr. Galbraith-Kent helped to guide us in directions that would make our study even more unique and was available to offer further guidance and/or help when asked.  Furthermore, Dr. Galbraith-Kent was available to assist us with getting the Berlese funnel set up and running to get our samples up and cooking.

Appendix A

Organism Density (Number of Organisms/40cm^2)
Springtail

19

Mite

6

Homoptera

3

Diptera

2

Small Black Beetle w/ textured back

2

White Larvae

2

Brown Spider

1

Gastropod

1

Rounded, Dark Brown Beetle

1

Small Spider w/ dark patterns

1

Winged Ant

1

Thrip

1

Dark Brown Striped Fly w/ spotted wings

1

Transparent catapillar esque

1

Protura – Black segmented conehead

1

Hairy horseface organism

1

Table One:  Soil Invertebrate Numbers and Organisms for Pioneer Park, Kenton County, Kentucky.

Organism Density (Number of Organisms/40cm^2)
Mite

16

Springtail

13

Short, round, brown springtail

7

Small, pink springtail

3

White Larvae

3

Isopod

2

Araneae

1

Psocoptera – white w/ red eyes

1

Large, tan/white larvae

1

Round, black beetle

1

Brown, textured beetle

1

Hymenoptera – ant

1

Psuedoscorpion

1

Catapillar (larvae)

1

Rounded, smooth, brown beetle

1

Table Two:  Soil Invertebrate Numbers and Organisms for Withrow Nature Preserve, Cincinnati, Ohio.

Pioneer Park Soil Invertebrate CommunityGraph One:  Rank Abundance Data Figure for Pioneer Park, Kenton County, Kentucky.

soil invertebrates

Graph Two:  Rank Abundance Data Figure for Withrow Nature Preserve, Cincinnati, Ohio.

Ordovician and Silurian Communities

Ordovician and Silurian Communities
By: Linda Zantout

Taxon Count Life Habit Complete or Broken Body or Mold Period
Trepostomes 2 Epifaunal Suspension Feeder Broken Body Ordovician
Platystropha 4 Epifaunal Suspension Feeder Complete Mold Ordovician
Homotrypa 1 Epifaunal Suspension Feeder Broken Mold Ordovician
Batostomella 3 Epifaunal Suspension Feeder Broke Mold Ordovician
Strophomena 1 Epifaunal Suspension Feeder Broken Mold Ordovician
Crinoids 3 Epifaunal Suspension Feeder Broken Body Ordovician
Laptaena 1 Epifaunal Suspension Feeder Broken Mold Ordovician

 

 

Data Analysis and Interpretation

 

1. Depending on the sample size being measured, different sets of individuals will be studied which may lead to different results in measuring species richness. Thus, results regarding species richness are most valid when the sampling sizes are for the most part equated. For example, within the class data we see 20 different types of brachiopods and 17 different types of bryozoa and thus a valid deduction would be that the species richness is higher in brachiopods. However, in a different sample in which the number of individual species in each phylum is added, there are 131 brachiopods and 20y bryozoa. In this sample set, it would be valid to say that the species richness of bryozoa is greater than that of brachiopods.

 

2.

Phylum Genera                           Count            Simpson Index
Bivalve Ambonychia

1

0.00212766

Bivalve Carotidens

1

0.00212766

Brachiopod Platystrophia

31

0.065957447

Brachiopod Lepidocyclus

16

0.034042553

Brachiopod Strophomena

14

0.029787234

Brachiopod Hypsiptycha

12

0.025531915

Brachiopod Thaerodonta

11

0.023404255

Brachiopod Hebertella

8

0.017021277

Brachiopod Zygospira

7

0.014893617

Brachiopod Megamyonia

5

0.010638298

Brachiopod Hesperorthis

4

0.008510638

Brachiopod Diceromyonia

3

0.006382979

Brachiopod Hiscobeccus

3

0.006382979

Brachiopod Rafinesquina

3

0.006382979

Brachiopod Retrisirostra

3

0.006382979

Brachiopod Glyptorthis

2

0.004255319

Brachiopod Leptaena

2

0.004255319

Brachiopod Rhyncotrema

2

0.004255319

Brachiopod Sowerbyella

2

0.004255319

Brachiopod Orbiculoidea

1

0.00212766

Brachiopod Pentamerus

1

0.00212766

Brachiopod Reserella

1

0.00212766

Bryozoa “trepostome”

83

0.176595745

Bryozoa Batostomella

36

0.076595745

Bryozoa Homotrypa

17

0.036170213

Bryozoa Dekayella

14

0.029787234

Bryozoa Prasapora

12

0.025531915

Bryozoa Rhombotrypa

8

0.017021277

Bryozoa Monticulipora

6

0.012765957

Bryozoa Parvohallopora

6

0.012765957

Bryozoa Dimesopora

5

0.010638298

Bryozoa Aspidopora

4

0.008510638

Bryozoa Fistulipora

4

0.008510638

Bryozoa Homotrypella

3

0.006382979

Bryozoa fenestrate

2

0.004255319

Bryozoa Heterotrypa

2

0.004255319

Bryozoa Rhimdictya

2

0.004255319

Bryozoa Dekayia

2

0.004255319

Bryozoa Batostoma

1

0.00212766

Conularid Cornulites

6

0.012765957

Coral Calapoecia

4

0.008510638

Coral Rugose coral – unid

3

0.006382979

Coral Favosites

1

0.00212766

Coral Grewingkia

1

0.00212766

Coral Protarea

1

0.00212766

Coral Streptalasma

3

0.006382979

Crinoid unidentified

75

0.159574468

Monoplacaphoran

1

0.00212766

Porifera Astylospongia

6

0.012765957

Porifera Hindia

3

0.006382979

Stromatoporoids

17

0.036170213

Tentaculitid Tentaculites

3

0.006382979

Trilobites

6

0.012765957

TOTAL

470

1

 

Taphonomy and Life Habits

1. The community seems to be heavily dominated by epifaunal suspension feeders.

2. The fossils can be divided into two sizes: small ones such as the trepostomes and other bryozoans and then bigger ones such as the brachiopods. However amongst each one of these divisions, the sizes are evenly distributed. Also, there are outliers, those that are significantly smaller or bigger, but that is to be expected.

3. What’s missing from the biota is more specimens of coral and porifera and some of cephalopods although they are generally harder to preserve so it makes sense.

4. The mode of preservation was generally mold or cast in that there was an imprint left behind in the rock in which the shape and texture of the shell could be seen. Other than this, there were some that had the body preserved but those were mainly bryozoans, which had hard body parts that were able to withstand the time and preservation.

5. Based on observations, it can be inferred that the organisms resided in a warm shallow marine environment and were deposited and preserved  at the bottom in sediments or mud after they died.

6. To a certain extent, the collection does represent the original living community. Of course however, there are a number of species missing due to the fact that they’re usually not preserved well or rare in general but there was a large percentage of brachiopods and bryozoans which is what mostly would have been making up the original living community.

7. The main difference between the Ordovician and Silurian communities is that the organisms of the Ordovician period were all marine organisms whereas the Silurian brought with it the appearance of the first few organisms able to start their way onto land. This makes sense since all the organisms we found were marine based and we were looking in Ordovician beds.

Measuring Respiration of Germinating and Non-germinating Peas

Measuring Respiration of Germinating and Nongerminating Peas
By: Krunal Patel

Introduction

          Living cells require transfusions of energy from outside sources to perform their many tasks – for example, assembling polymers, pumping substances across membranes, moving, and reproducing (Campbell, and Reece 162). Heterotrophs obtains its energy for its cells by eating plants that makes it own food (Autotrophs); some animals feed on other organisms that eat plants. The most beneficial catabolic pathway in an organism is cellular respiration, in which oxygen and glucose are consumed and where carbon and water become the waste products. The purpose of cellular respiration is to convert glucose into ATP(energy) for the organism. Respiration consists of glycolysis, the Krebs Cycle, and the oxidative  phosphorylation. Glycolysis, which occurs in the cytosol, breaks the six carbon glucose molecule into two pyruvates. During this stage two ATP and two NADH molecules are made. The next step in respiration is the Krebs cycle. The Krebs cycle uses the two pyruvates made during glycolysis and converts them to Acetyl-CoA and carbon dioxide to make three NADH, one FADH2,  and two CO2 through redox reactions, and goes to the Electron Transport Chain. ATP is also formed during the Krebs cycle (Campbell, and Reece 166). Since two pyruvates are made during glycolysis, the Krebs cycle repeats two times to produce four CO2, six NADH, two FADH2, and two ATP (Campbell, and Reese 166). The last stage in cellular respiration is the Oxidative phosphorylation Electron Transport. The Oxidative phosphorylation occurs in the inner membrane of the mitochondria. The electron transport chain is powered by electrons from electron carrier molecules NADH and FADH2 (Campbell, and Reese 166). As the electrons flow through the electron chain, the loss of energy by the electrons is used to power the pumping of electrons across the inner membrane. At the end of the electron transport chain, the electrons from the inner membrane bind to two flowing hydrogen ions to form water molecules. The protons, outside the inner membrane, flow down the ATP gradient and make a total of thirty two ATP (Campbell, and Reese 166).

In this experiment, an apparatus called a respirometer is used. A respirometer is a tool used to observe exactly how much oxygen was consumed by the peas and the glass beads. Since the carbon dioxide produced is removed by reaction with potassium hydroxide (Forming K2CO3 + H2O as shown below), as oxygen is used by cellular respiration the volume of gas in the respirometer will decrease. As the volume of gas decreases, water will move into the pipet. This decrease of volume, as read from the scale printed on the pipet, will be measured as the rate of cellular respiration (Cell Respiration).

CO2 + 2KOH —> K2CO3 +H2O

The purpose of this lab was to measure the rate of cellular respiration. There are three ways to measure the rate of cellular respiration. These three ways are by measuring the consumption of oxygen gas, by measuring the production of carbon dioxide, or by measuring the release of energy during cellular respiration (Respiration). In order to measure the gases, the general gas law must be understood. The general gas law state: PV=nRT where P is the pressure of the gas, V is the volume of the gas, n is the number of molecules of gas, R is the gas constant, and T is the temperature of the gas (Respiration). The rate of respiration of germinating and non-germinating peas in this experiment was determined by the consumption of oxygen. Potassium Hydroxide (KOH) was used to alter the equilibrium. KOH removed the carbon dioxide and oxygen was used by cellular respiration thus decreasing the gas in the respirometer. The rate of respiration in germinating peas was compared to the rate of the non-geminating peas. These peas were placed in two different temperatures: 10ºC and 23ºC.

The hypothesis of this lab states that if the peas are germinated then the rate of cellular respiration will be higher in both room temperature and cold temperature. If the temperature of water is cooler than room temperature, then the process of cellular respiration of the peas will decline.

Materials

v  Room-Temperature Water Bath                      Nonabsorbent Cotton

v  Cold Water Bath                                             15% Potassium Hydroxide (KOH) Solution

v  Container of Ice                                              Dropping Pipets

v  Paper (White or Lined)                                   Forceps

v  Water                                                              Thermometers

v  Germinating Peas                                            Stopwatch (Timer or Clock)

v  Nongerminating Peas                                      Calculators (Optional)

v  Glass Beads                                                    Absorbent Cotton Balls

v  Respirometers                                                 Graduated Tube

Procedure

Setup of Respirometers and Water Baths

There are two water baths (trays of water) to buffer the respirometers against temperature change and to provide two temperatures for testing: room temperature and a colder temperature (Approx. 10°C). Place of sheet of paper in the bottom of each water bath. This will make the graduated pipet easier to read. Next, place a thermometer in each tray. If necessary, add ice to the cold-temperature tray to further cool the water to get it as close to 10°C as possible. While waiting for the cold- water temperature to stabilize at 10°C, prepare the three respirometers to test at room temperature, and prepare an identical set of three respirometers to test at the colder temperature.

 

 

Prepare Peas and Glass Beads

Respirometer 1: Put 25 mL of H2O in your 50-mL graduated plastic tube. Drop in 25 germinating peas. Determine the volume of water that is displaced (equivalent to the volume of peas). Record the volume of the 25 germinating peas. Remove these peas and place them on a paper towel.

Respirometer 2: Refill the graduated tube to 25 mL with H2O. Drop 25 dry, nongerminating peas into the graduated cylinder. Next, add enough glass beads to equal the volume of the germinating peas. Remove the nongerminating peas and beads and place them on a paper towel.

Respirometer 3: Refill the graduated tube to 25 mL with of H2O. Add enough glass beads to equal the volume of the germinating peas. Remove these beads and place them on a paper towel.

The independent variable is the type of peas (Germinated or Nongerminated) and the temperature (Room or Cold Temperature). The dependent variable is the consumption of oxygen from all 6 respirometers. The control group is respirometer three from both temperatures that consists of only glass beads.

Respirometer Assembly

This requires three respirometers for room-temperature testing and three respirometers for cold-temperature testing.

To assemble a respirometer, place an absorbent cotton ball in the bottom of each respirometer vial. Use a dropping pipet to saturate the cotton with 2 mL of 15% KOH. (Caution: Avoid skin contact with KOH. Be certain that the respirometer vials are dry on the inside. Do not get KOH on the sides of the respirometer.) Place a small wad of dry, nonabsorbent cotton on top of the KOH- soaked absorbent cotton. The nonabsorbent cotton will prevent the KOH solution from contacting the peas. It is important that the amount of cotton and KOH solution be the same for each respirometer.

  • Place 25 germinating peas in the respirometer vial(s) 1.
  • Place 25 dry peas and beads in your respirometer vial(s) 2.
  • Place beads only in your respirometer vial(s) 3.

Insert stopper fitted with a calibrated pipet into each respirometer vial. The stopper must fit tightly. If the respirometers leak during the experiment, you will have to start over.

Placement of Respirometers in Water Baths

Place a set of respirometers (1, 2, and 3) in each water bath with their pipet tips resting on lip of the tray. Wait five minutes before proceeding. This is to allow time for the respirometers to reach thermal equilibrium with the water. If any of the respirometers begins to fill with water, the experiment will have to restarted.

After the equilibrium period, immerse all respirometers (including pipet tips) in the water bath. Position the respirometers so that it’s easy to read the scales on the pipets. The paper should be under the pipets to make reading them easier. Do not put anything else into the water bath or take anything out until all readings have been completed.

Take Readings

Allow the respirometers to equilibrate for another five minutes. Then, observe the initial volume reading on the scale to the nearest 0.01 mL. Record the data in Table 1 for Time 0. Also, observe and record the temperature. Repeat your observations and record them every five minutes for 20 minutes.

(Cell Respiration)

Results/Data Collection

Table 1: Respiration of Peas at Room Temperature

Respirometer 1 Germinating Peas

Respirometer 2 Dry Peas + Beads

°C

Time (Min)

V of Pipet

ΔV

Corrected ΔV

V of Pipet

ΔV

Corrected ΔV

26

0

.81

-

-

.95

-

-

26

5

.71

.1

.06

.92

.03

.015

26

10

.66

.15

.105

.91

.04

.015

26

15

.59

.22

.17

.9

.05

.015

26

20

.52

.29

.23

.89

.06

.015

*All values are in mL except °C and Time

 

Table 2: Respiration of Peas at 10° C

Respirometer 1 Germinating Peas

Respirometer 2 Dry Peas + Beads

°C

Time (Min)

V of Pipet

ΔV

Corrected ΔV

V of Pipet

ΔV

Corrected ΔV

10

0

.2

-

-

.20

-

-

10

5

.31

.11

.035

.28

.08

.02

10

10

.4

.20

.12

.29

.09

.02

10

15

.43

.23

.18

.26

.06

.025

10

20

.45

.25

.20

.26

.06

.01

*All values are in mL °C and Time

Respiration of Peas

 

Discussion/Conclusion

The results of this lab show that the germinating peas had consumed more oxygen at a faster rate than the non-germinating peas and the beads had. The non-germinating peas and the beads showed to consume barely any oxygen at all. In this lab, the germinating peas respiration rate proved to be faster than the respiration rate of non-germinating peas. Finally, this experiment showed that respiration rates increase as the temperature increases. Concluding that temperature and respiration rates are directly proportional and have a direct relationship to each other. From this experiment, it can also be concluded that the germinating peas that were undergoing the process of cellular respiration had a much higher oxygen consumption rate than the consumption rate in non-germinated peas and the glass beads. The non-germinating peas shows hardly any consumption of oxygen. Since the germinating peas are germinating or sprouting, they require a more extensive amount of energy or ATP. This allows them to have high oxygen consumption rates or respiration rates in this experiment. In addition to the germinating peas, the non-germinating peas, are not germinating so because of this they do not need significant amount of ATP production. Therefore, the non-germinating peas have a significantly low rate of respiration in comparison with the germinating peas. The rate at which they respire was most prevalent in the first respirometer since they were all germinating peas. The data table and graph accurately depict this idea or trend with the germinating and non-germinating peas.

Numerous errors could have occurred during the lab. Miscalculation of the glass beads which went into the respirometer, and hence would ruin the controlled results. The seals on the respirators may not have been completely air-tight which may have caused a leak and therefore oxygen would have been lost altering the data.  The temperature may have been slightly off in the water baths. There was also the problem of reading the scales on the pipets which could have lead to improper measurements of the water position.

To improve this experiment, more accurate and precise instruments can be used such as a advanced pipet which has scales that are easy to read. Also a completely air tight respirometer instead of using petroleum jelly which water can still leak in.

 

Literature Cited

Campbell, Neil A., and Jane B. Reece. Biology. Eighth Ed. San Francisco: Pearson Benjamin Cummings, 2008. Print.

Cell Respiration. AP Biology Laboratory 5: Carolina Biological Supply Co., 2005. Print.

“Respiration.” StudyMode.com. StudyMode.com, 06 2011. Web. 06 2011. <http://www.studymode.com/essays/Respiration-713319.html>.