DROSOPHILcross over

INTRODUCTION

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Laboratory Exercise Guidelines Please Note: You must type your lab report and submit it on HUSKYCT via “SafeAssign.” This program is designed to scan your document for plagiarism against your peers (past and current), journal articles, and other online sources. Please use size 12 font, double-space formatting, and limit it to no more than 4 pages. Hardcopies will not be accepted under any circumstances.

The majority of the lab reports, which are collected and graded in this course, are graded as follows:

Introduction (10%) Relevant background information The major aim(s) of the experiment

Theoretical Questions (30%) Answer any provided questions Offer expected results (hypothesis)

Results (30%) Answer any provided questions Provide clear and detailed data Summary of results Any figures, tables, and graph axes are properly labeled

Discussion/Conclusion (30%) Answer any provided questions Summarize experiment Do your data support the hypothesis? Describe errors and possible sources of errors Concluding remarks (significance of results)

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MCB 3413: CONCEPTS OF GENETIC ANALYSIS

Fly Pushing A number of laboratory exercises in this course require the use of Drosophila melanogaster (commonly referred to as fruit flies) as a genetic model. The following section offers some guidelines for working with the flies.

Figure 0.1 Fly workstation example (laboratory classroom set may differ)

I. Fly Anesth etization 1. Unclamp the tube, which connects to the etherizer bottle 2. Bump the bottle gently on the bench top to knock the flies to the bottom of the bottle 3. Quickly remove the bottle top and invert the bottle onto the etherizer bottle 4. Tap the flies out (take care not to tap too hard—fly food may enter the etherizer) 5. Upon removal of the flies from the bottle, replace the bottle top and return the bottle to the tray 6. Unclamp the tube, which connects to the CO2 pad that is located on the dissecting scope 7. Gently empty the flies from the etherizer onto the pad; flies are now anesthetized and ready for

manipulation *when finished, be sure to re-clamp the tubes that supply CO2 to the etherizer and pad*

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INTRODUCTION

II. Identif yi n g S exes an d Gen et ic Markers

S e x :

Using a fine-bristle brush, manipulate the flies in a manner that will enable you to view the posteriors of the flies. Familiarize yourself with sexing the flies in order to set up your fly crosses. Virgin females will be used in your cross set-ups. Your TA will explain the significance of using virgins to set up a genetic cross.

Figure 0.2 Characteristics of adult Drosophila males and females. Males have a dark posterior region (more obvious in older males) and a brownish ring surrounding the genital plate (Panel a). Females lack the genital plate ring and are slightly larger than males. Males also contain sex combs, which are bristles located on the forelimbs (Panel b).

G e n e t i c M a r k e r s :

Table 1 summarizes the recessive and dominant mutations that are used as markers for analysis of meiotic chromosome segregation. The figure on the following page shows different genetic markers and the associated phenotype. Manipulate the flies in a manner that will enable you to view their different anatomical features. Familiarize yourself with these phenotypes.

Table 0.1

Mutation Location Type of Mutation Phenotype w (white) X recessive white eyes y (yellow) X recessive yellow body Cy (Curly) 2 dominant curled wings

Sco (Scutoid) 2 dominant loss of bristles, scutellars Sb (Stubble) 3 dominant short, thin bristles

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MCB 3413: CONCEPTS OF GENETIC ANALYSIS

Figure 0.3 Mutations used as genetic markers to monitor chromosome segregation in meiosis. For further details, see Table 0.1.

III. Cross S et-u p s

All crosses should consist of 4–5 female virgins and 4–5 males. Record genotypes (if known) and phenotypes of flies used. Use a fresh food vial for each cross and label it with your name, section number, and cross identification. Leave vial on its side until all flies have woken up.

Suggestion: In your notebook, record the date on which the cross was set-up.

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USE OF MICROPIPETTORS Micropipettors are designed to deliver from 1 to 1000µl (0.001-1.0ml) with extreme accuracy within the specified volume range. These pipettors must always be used with a disposable tip. It is very important to remember that different types of tips are not always interchangeable between different sizes or brands of pipettors.

Micropipettors are sensitive and expensive lab tools! Please review these instructions before each use.

1. Check the range of the pipettor. It is displayed at the top of the pipettor. You will be using pipettors having ranges of 1-10µl, 10-100µl, and 100-1000µl. Do not force the pipettor beyond its designated range - you will damage the pipettor.

2. Set the desired volume by unlocking and then turning the volume adjustment knob until the correct volume appears on the digital indicator. Then re-lock the knob prior to usage. For best results always approach the desired volume by dialing down from a larger volume setting.

3. Attach a new disposable tip to the shaft of the pipettor. Press the tip on firmly, using a slight twisting motion, to ensure a positive, airtight seal. Use WHITE tips for the 1-10µl pipettor, YELLOW tips for the 10-100µl pipettor, and BLUE tips for the 100-1000µl pipettor.

4. Depress the plunger to the FIRST positive stop. This is the volume displayed on the digital indicator.

5. Holding the pipettor vertically, immerse the lower third of the tip into the sample liquid, and allow the plunger to return SLOWLY to the UP position. Do not allow it to snap up!

6. Wait 1-2 seconds to ensure that the full sample volume has been drawn into the tip.

7. Withdraw the tip from the solution. Should any liquid remain on the outside of the tip, wipe it off carefully with a Kimwipe®. (Do not touch the tip opening!)

8. To dispense the sample, place the tip end against the inside wall of the tube, flask, etc. and depress the plunger to the FIRST stop. Wait several seconds, then depress to the SECOND stop (the bottom of the stroke) to expel any residual liquid in the tip.

9. Keeping the plunger depressed, withdraw the tip from the tube, and then allow the plunger to return slowly to the UP position. Discard the tip into the sharps container by pushing the ejector button.

INTRODUCTION

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MCB 2610: FUNDAMENTALS OF MICROBIOLOGY LAB MANUAL

LABORATORY NOTEBOOK SUGGESTIONS 1. THREE-RING BINDER AS A LAB NOTEBOOK Keep your lab notes in a three-ring binder with extra paper available for recording observations.

2. LABEL EACH NOTEBOOK ENTRY The lab notebook should be a well organized, easy to use compilation of your procedures,

observations, and results. It is not necessary to repeat the details of procedures already in your lab manual, but you should always indicate by a suitable label, the conditions under which a procedure was applied immediately above or adjacent to any list of results.

3. ORGANIZE YOUR NOTEBOOK CAREFULLY The lab notebook is a scientist’s primary source of laboratory experimental procedures and

data collected. Memory fades quickly, hence results should always be written down at the time they are observed, along with any other significant factors. You should keep a complete, well organized record of your observations. You will use this data for diagnosing unknowns later in the course.

4. ANSWER QUESTIONS ABOUT LAB EXERCISES In addition to results, most exercises include questions and provide a space for discussion/

conclusions. These should be answered at the end of the write up for that exercise and handed in when requested by your lab instructor in the format requested. In writing up each lab, answer the question, “What did this exercise show?” Be sure to explain any unusual, contradictory, or inconclusive results.

5. KEEP ALL WRITE-UPS FOR EACH EXERCISE IN ONE PLACE All the results of a given exercise should be written up in one place even if the data are collected

over several lab periods.