profile module: physiology of bacteria ag cypionka · profile module 'physiology of...

Profile Module: Physiology of bacteria

AG Cypionka

30.11 –11.12, 2009 C o n t e n t s Page Program .......................................... 2 Safety hints ..................................... 2 How to write the report .................. 3 Seminar presentations .................... 3 Exp. I Isolation and counting ....... 4 Exp. G Growth .............................. 12 Exp. R Aerobic Respiration .......... 17 Exp. P Proton translocation ......... 20 Exp. T Transport ........................... 23 Schedule Group A B C Mo I I I Tu P ↓ T ↓ R ↓ We P ↓ T ↓ R ↓ Thu R ↓ G ↓ P ↓ Fr R ↓ G ↓ P ↓ Mo T ↓ R ↓ G ↓ Tu T ↓ R ↓ G ↓ We G ↓ P ↓ T ↓ Thu G P T Fr P r e s e n t a t i o n s o f R e s u l t s Experiment: Supervised by: G = Growth Anette Schulte/Michael Pilzen T = Transport Michael Pilzen R = Respiration Anette Schulte P = Phage Tim Engelhardt/ Wiebke Landwehr I = Isolation / Counting of soil bacteria Anette Schulte/Michael Pilzen

Thanks to Daniela Wischer, Odeta Shuti, Johannes Holert and Sonja Standfest for the English translation.

Profile module 'Physiology of Bacteria' AG Cypionka - 2 -

P R O G R A M Theory: Start every morning at 8.30 (until ≈10) in Seminar room W15-0-023 1. Week: Theory of the experiments Mo Group organisation, prepare media and agar plates afternoon: Exp. I Late afternoon: "How to give an oral presentation" and "How to write a report" Tu Introduction and incubation Exp. T, Continue Exp. I, Start Exp. G/R/T/P, Introduction to microscope usage We Introduction growth, Exp. G Thu Introduction Phages, Exp. P Fr Introduction respiation Exp. R, 2. Week: Seminar and results presentation Mo – We: Oral Presentations (max. 10 minutes + 5 min discussion) Fr: Final Assessments presented by the students Practice (see schedule on page 1) • Max. 16 students in four groups (A, B, C, D), resp. 8 groups of two students (A1, A2, B1, ...) • 5 Experiments, I (running 10 days), G, R, P und T (2 days, round Robin) • Every experiment is performed twice (A1 and A2 ...) S a f e t y i n t h e L a b o r a t o r y • Please wear a laboratory coat! Gloves and Peleus balls for pipetting are provided. Do not eat, drink and smoke in the laboratory! Keep your desk clean and put clear labels on all vessels. • We are not working with pathogens. However, used plates are collected and autoclaved separately. Make sure that no fungal spores are released! • Toxic compounds are collected and disposed in special containers! • Do not use any laboratory equipment unless you are instructed about its use!


How to write the report

Please use a bound laboratory journal to record the everyday work step by step. Also write down the main points of the introductory lectures and the evaluations after the experiments. These notes are useful for the introduction of your report and for answering the questions.

In addition to the seminar presentation, the final report is required for passing the course. It must be written by everyone independently. (If reports show suspicious similarities, the concerned persons will be invited to a written test!) Write the report in such a way that one can understand - even after several years - what was the sense and outcome of the experiments. Write a short introduction that explains the principle of the experiment and your expectations. Do not repeat the instructions, describe methods only when you used different from those in the instructions. Tables and diagrams can be copied for a small group. Those data are not results – results are answers to your questions! Please explain the steps of your calculations with the help of a labelled curve where possible. Discuss possible differences between results and your expectations. The questions listed in the instructions preferably should be answered in the context of your introduction or discussion. The report should be compiled with the help of a word-processing program. For each experiment, the report should contain 2-3 pages of written text (font size 12, single-spaced) in addition to tables and diagrams. For writing the report read the guide for creation of scientific texts (www.icbm.de/pmbio). Please print out the report and staple together each experiment individually with your name written on it. In addition, create one pdf file of the report and sent it to [email protected] (Recommendation: Freeware PDF-Creator). Report deadline For the verification of a successful participation, the report must be handed in until end of January (P.O. Box, Cypionka, staircase of the ICBM building). Please collect the reports during the office hours (Friday, 10 -12 h) after the announcement that they have been read. Seminar Presentations While during the first week the theory is closely related to the practical program, in the second week, seminar presentations of current research topics are given by the participating students. You can use Powerpoint for this. Please do not show copies from articles, but create few (!) own slides showing the important points. An introduction to the contents of the article is much more important than a complete summary! What should the listeners learn about the subject? Speak for 10 min, plan 5 min for the discussion. The supervisors can help if you have questions during the preparation. Obligatory attendance Attendance is not only compulsory for the practical part, but also for the seminars. Absence without valid excuse, will lead to loss of the course certification.


Experiment I = Isolation and counting of soil bacteria Background In this experiment bacteria are isolated in pure culture either directly or after previous enrichment. Every group varies medium or incubation parameters in order to study their influence on the outcome of the enrichment. Enrichments under different conditions and direct isolation of bacteria are expected to give different results. (Do no forget to explain your expectations.) Principle of the experiment In the first part, the influence of different conditions on the outcome of enrichments is studied. The basic experiment is the enrichment and isolation of bacteria which grow aerobically at room temperature and low salinity with lactate as substrate and ammonia as N source. Parameters such as temperature (~20 °C, 37 °C, pasteurisation of the sample at 80 °C for 10 min), N source (NH4

+, N2), pH (acidic or basic) and salt concentration (0 or 3 %) are varied. Pure cultures are obtained (from subcultures) by the 13-streak technique on agar plates. In the second part, direct isolation and counting of the live cell number (based on serial dilutions) is performed. Isolates are compared with those obtained after enrichment (basic experiment from first part). Time schedule The experiment will last ten days. Its progress will be determined by the growth of bacteria. A rough scheme is given here: Day 1. • Preparing and autoclaving the media • Pouring the agar plates • Inoculation of the enrichment cultures • Starting counts and direct isolation: Prepare serial dilutions from the soil samples and plate them on agar medium that will be provided by the course staff. Day 2. • Checking the turbidity of the enrichments (write down your observations in your table), if they are already turbid, proceed as described under day 4. Day 3. Visual checking (main experiment day for the other experiments) Day 4. • Check culture microscopically, make a microphotograph, streaking out subcultures and colonies from the direct isolation on agar plates. Day 5. Visual checking (main experiment day for the other experiments)


Day 8. • Microscopy, second dilution streak on agar plates . Day 9 and 10. • Description of colonies and single cells (microphotography!) • Gram test, oxidase test, catalase test Required materials: • Cylinder, Erlenmeyer flasks (each 250 ml), magnetic stirrer • Distilled water, ingredients for the media, scale • pH-meter, calibration buffer, 1 M HCl • Erlenmeyer flasks, 100 ml, cotton plug, aluminium foil • Screw top bottles (50 ml) • Screw top tubes for the serial dilution • Drigalski spatula • Autoclave (coordinate use with other groups!) • Bunsen burner, inoculation loop • Sterile pipettes, 5 ml und 1 ml • Phase-contrast microscope with digital camera • Stereo microscope Preparing the medium For each group: • 4 x 100-ml Erlenmeyer flasks with 20 ml medium • 12 Agar plates, each containing 25 ml medium (basic recipe and variations) Lactate medium: basic recipe Components Per liter medium Concentration Distilled water 950 ml 55.5 M (NH4)2SO4, mw 132.14 1.06 g 8 mM MgSO4 * 7 H2O, mw 246.48 0.10 g 0.4 mM CaCl2 * 2 H2O, mw 147.02 0.03 g 0.2 mM Sodium lactate solution (50% = 5.71 M ) 3.5 ml 20 mM Trace element solution SL9 1 ml (for composition see experiment G) Prepaire and autoclave separately in 100-ml bottle: Potassium phosphate buffer, pH 7.0, 1.0 M 50 ml 50 mM !! Pay attention to the variations, not everything belongs to each medium !! Dissolve the components with distilled water in a graduated cylinder, set the pH to 7.0 and fill it up with water to the end volume. Give 20 ml of each medium into 100 ml Erlenmeyer flasks, close them with cotton plug and put them into the autoclave. After cooling down add 1ml Potassium phosphate buffer (autoclaved separately) to 20 ml medium


Agar medium The same medium can be mixed with 1.5 g agar per 100 ml and then put into the autoclave. After cooling down to 60°C add 5ml Potassium phosphate buffer (autoclaved separately) to 100 ml medium and pour the medium into sterile Petri dishes. Protect the plates from drying out! !Immediately rinse dirty agar vessels with hot water and brush! After use, put the plates into plastic bags. Make sure that no fungal spores are released! Enrichment and Isolation (basic experiment and variation) I-0: Basic recipe (also used for counts and direct isloation) • This is needed for each group as everybody will inoculate I-0 as control; prepare enough sterile medium in flasks and agar plates for all the groups (500 ml liquid medium, 2x500ml agar medium) Variations I-A: Increased temperature • Use basic recipe, but incubation occurs at 37 °C I-B: Pasteurisation • Use basic recipe, but pasteurise the inoculum (1 ml) before inoculation of the enrichment. To do this, transfer the culture into a tightly closed test tube, and heat this, completely submerged, at 80 °C for 10 min. Incubation of the enrichment occurs at room temperature. I-C: N 2 as nitrogen source • Basic recipe without ammonium sulfate I-D: Salt • Basic recipe, in addition 3.0 g NaCl/100 ml I-E: Acidic pH • Basic recipe, pH adjusted to 6;instead of potassium phosphate buffer use 30mM acetate buffer,ph 6 (will be provided) I-F: Alkaline pH • Basic recipe, pH adjusted to 8;instead of potassium phosphate buffer use 30mM Tris buffer,ph 8 (will be provided) Course of the experiment • Enrichment in the liquid culture→ • subculture from that (1st culture obtained) → • streak out plates from that (13-streak method) → • streak out a 2nd time → • get 2 "pure cultures" of that → • Gram-, catalase and oxidase tests of pure cultures. Inoculation of the enrichment The garden soil’s material for the enrichments should be the same for all groups, so that each of them can explain the different results under the enrichment’s conditions. Take a little bit from the soil material, dilute it in the 10-fold volume of tap water, let particles settle down for 5 min. Pour


the upper liquid of the mixture into another test tube. From this, each group uses 1 ml for the inoculation of the enriched cultures. All cultures, apart from the I-A variation, will be incubated in the lab, at room temperature (appr. 20 - 25°C), they will be daily checked and mixed. Subculture As soon as an enrichment culture becomes turbid, check it under the microscope (draw / photograph the cell types for the report). Make a subculture (inoculate with 1 ml, incubate under the same conditions), continue to check daily. Dilution streak on agar As soon as one culture is grown, compare the cell types with those of the first culture (phase contrast microscope), and make a dilution streak (13-streak method). Check daily, record the appearence of colonies, describe the colony types (binocular microscope). Second dilution streak Make a second dilution streak from each characteristic cell type and win “pure cultures” from that and for further characterisation.

Counting and direct isolation Course of the experiment Serial dilution 6 x (1 + 9) → • inoculate from that 5 plates with 0.1 ml (= 10-7 - 10-3 g soil) → • 2 plates from the plates with nice colonies (13-st.-m.) → • 2 „pure cultures“ from that → • perform with those Gram test etc. For the direct isolation and count, 6 screw top tubes with 9 ml tap water each, will be autoclaved. After cooling , 1g soil will be transferred into the first screw top tube and mixed thoroughly for 3 min. Now, to make a serial dilution, 1 ml of the above mixture will be taken out with a sterile pipette and transferred into another test tube (nr. 2), and mixed. In each case use sterile pipettes! From the test tubes nr 6 to nr 2 (in respective order), 0.1ml will be taken out of each with a sterile pipette and will be plated out on agar (basic recipe) with a sterile Drigalski-

spatula, ( this corresponds to 10-7 - 10-3 g soil). No parallel attempts are necessary. Instead of that, the results of all groups will be compiled for the evaluation. The agar plates will be incubated at room temperature and daily checked. Compile tables with the numbers of colonies every day. Isolate in pure cultures the most frequent or the most interesting lactate utilizers (from the highest dilution degrees) (second streak according to the enrichment culture). Characterisation of “pure" cultures The following tests should give a hint on the physiological diversity of the isolates, they are not aiming at a taxonomic classification of the isolates (but think about a possible classification in your report). Gram test This is no real Gram staining, but gives the same result in 90 % of all cases within a minute. Reagent: 3% KOH (will be provided)


Procedure: Give one drop of the 3% KOH solution onto a slide. Cell material from colonies or from centrifuged cells is mixed with this drop for 3 to 5 s with an inoculation loop. If slimy filaments are visible when the inoculation loop is carefully lifted, the cells are Gram-negative. With Gram-positive cells no slimy material will be formed. Literature : Gregersen, T. Rapid method for distinction of Gram-negative from Gram-positive bacteria. Eur. J. Appl. Microbiol. Biotechnol. 5:123-127 Oxidase test Reagents: H2O 100 ml Ascorbic acid 0.1 g Tetramethyl-p-phenyldiamin-HCl 1.0 g Attention, carcinogen! Reagents (will be provided, freshly prepared every day!). Do not touch by hand and do not spill! After using, wash immediately the vessels and pipettes! Procedure: Soak a strip of filter paper (cellulose) with some drops of the reagent (not too wet!). Give some bacterial mass taken from a colony with an inoculation loop on the strip, and rub it in by a rounded glass rod. Blue coloration shows oxidase activity. As a control compare with a preparation of strains of known acitivity. Catalase test Reagent: (will be provided) 5% H2O2 solution, produced by diluting a 30% stock solution with distilled water. Store in a cool and dark place. Do not touch by hand! Contamination (e.g. dust, metal etc.) provokes corrosion. Procedure: Place some bacterial mass from the middle of one fresh colony on a clean slide. Then give one drop of the diluted H2O2 solution onto it using a clean Pasteur-pipette. Formation of gas (O2) shows catalase activity. Evaluation Compile tables and register the daily observations and your procedures. In the report, describe with words the process of the enrichment. Calculate the number of the cultivated lactate-utilising bacteria per g soil. Compare the obtained pure cultures from the enrichments and direct isolations. Compare the main experiment and your variation. Give a physiological explanation for the observations of the other groups. List all the observed characteristics of the pure cultures and try a rough taxonomic classification. Compare the results with your expectations!


Calculation number of bacterial colonies in and on agar plates with different degrees of dilution (based on L. Cavalli-Sforza, Biometrie, Grundzüge biologischer Statistik, G. Fischer Verlag, 1972) ΣiCi x = ——— Σi(ni*zi) x = average cell number in the inoculated enrichment C1, C2, ... , Ci = number of individual counted colonies n1, n2, ... , ni = plates / degree dilutions z1, z2, ... , zi = dilutions Example:

Dilution: 10-6: 303, 290, 285 Colonies

Dilution: 10-7: 32, 21 Colonies

Average cell number x = 931/(3*10-6 + 2*10-7) = 2.91*108

Suggestion for Tables of the report: Table 1: Enrichment Group: Varied Parameter: Date Day Basic parameters Variation No. . . Observations / Action Observations / Action . . . . . .


Tables 2 and 3 are required already at the last day of the course! Table 2: Characterisation of pure cultures GROUP: From Enrichment: Basic conditions Variation:_________ Shape Motil. Gram Cat. Oxid. Shape Motil. Gram Cat. Oxid. From direct isolation: Basic conditions Shape Motil. Gram Cat. Oxid. Table 3: Direct counts of colony-forming bacteria Colonies on plates with g soil nach 10-3 10-4 10-5 10-6 10-7 Total count per g soil: 1 d (We) 2 d (Thu) 3 d (Fr) 6 d (Mo) 7 d (Tu) 8 d (We) 9 d (Thu)


E x p e r i m e n t G = Growth Aim The aim of this experiment is to obtain growth parameters of a fast-growing bacterium. These parameters will be the basis for further considerations of the course. In addition to the growth rate, which is determined on the basis of cell number and turbidity, the energy metabolism during fermentation and respiration is compared. This is done by determining the yield coefficients with and without oxygen. Approach Cells are incubated with shaking aerobically, as well as in closed tubes without O2. From aerobic flasks, samples are analysed every hour (or every 30 min). From the anaerobic cultures in the same medium, only the OD is measured. In a separate test, the dry mass per OD and per cell number is determined. Time schedule Day 1 • Preparation and autoclaving of media • Sterilisation of pipettes, tubes etc. • Cleaning of weighing dishes • Setting up precultures Day 2 • Inoculation of the studied cultures early in the morning • taking samples throughout the day, counting cells under the microscope • immediate plotting of data (linear and semilogarithmic) • Preparation of dry-mass determination Day 3 (in addition to new experiment!) • Determination of dry mass • Evaluation Materials needed For medium preparation: • Graduated cylinder (500 ml), Erlenmeyer flasks (500 ml), magnetic stirrer • Ddistilled water, media ingredients, scale • Potassium phosphate buffer, 1 M, pH 7.0 • Trace element solution SL-9 • Multivitamin solutionVL-7 • pH-meter, standard buffer, 1 M HCl • Baffled flasks (500 ml), felt cloth, tin foil for covering • Screw-cap tubes (22 ml) • Screw-cap bottles (50 ml) • Autoclave (co-ordinate usage with other groups!) • Bunsenburner, inoculation loop • Stock culture of the bacterial strain to be analysed • Shaking incubator, 30 or 37 °C


For dry mass determination: • Two light glass beakers (5 ml) • Tweezers, Pasteur pipettes • Refrigerated centrifuge with rotor for 250 ml beakers (coordinate usage!) • 2 centrifuge beakers (250 ml) with lids • Ammonium acetate buffer, ≈50 mM, pH ≈6.5, 1 l, keep closed! For growth experiment: • Sterile pipettes, 5 ml, 2ml • Photometer: Bausch & Lomb Spectronic 70 • Semi-micro cuvettes, preferably glass • phase-contrast microscope • Counting cell chamber with special cover glass • For motile cells: Lugol's solution (1 g J2 + 0.5 g KJ in 100 ml), use 1 + 1 dilution • Microliter syringe (50 µl) Preparing the medium: Each group needs: 2 * 125 ml in baffeld flasks for dry mass determination Two baffeld flasks with 100 ml medium for aerobic preculture and aerobic growth experiment Two screw-cap tubes (first in 50 ml bottles) for anaerobic preculture and experiment Glucose- Medium [Eppstein W, Kim BS (1971) J. Bacteriol. 108:639-644] per l Medium Concentration A) Mineral salts: Dist. water 930 ml 55.5 M (NH4)2SO4, MM 132.14 1.06 g 8 mM MgSO4, MM 246.48 0.10 g 0.4 mM CaCl2 * 2 H2O, MM 147. 0.03 g 0.2 mM A1)K2HPO4 buffer, pH 7.0, 1.0 M 70 ml (will be provided) 70 mM B) Glucose solution: D-Glucose, MM 180, 1.80 g/10 ml 5 ml 5 mM C) Multivitamin solution VL-7 1 ml (will be provided) D) Trace element solution SL-9 1ml (will be provided) To prepare 500 ml of solution A), dissolve the ingredients with distilled water in a graduated cylinder and fill up to final volume. Using another graduated cylinder (100 ml), transfer 99 ml of the solution into baffeld flasks for each anaerobic experiment; and 25 ml into 50 ml screw-top bottles for each aerobic preparation. Close the baffeld flasks with felt, tinfoil and elastic band; tightly screw the (half-full) screw top bottles. Transfer 50 ml from solution A1) in a 100 ml bottle. From solution B) transfer 10 ml into a screw-top bottle (50 ml) and firmly close this. Place all closed containers in a polypropylene beaker and cover the beaker with felt and tin foil


as protective shield. Solutions A) A1) B) are autoclaved for 20 min at 121 °C. Once the media have cooled to room temperature (taking special care of the closed vessels), complete the media by adding solution A1)( 7 ml per 100ml preparation, 1,75 ml per tube) glucose (1ml per 100ml, 0.25 ml per tube) as well as solution C and D (will be provided). For the anaerobic experiments, the medium is completed before it is transferred to a tube. Trace element solution SL-9 (will be provided) Distilled water ad 1000 ml Nitrilotriacetic acid (NTA) 12.8 g FeCl2 * 4 H2O 1.5 g CoCl2 * 6 H2O 190 mg MnCl2 * 2 H2O 100 mg ZnCl2 70 mg NiCl2 * 6 H2O 24 mg Na2MoO4 * 2 H2O 36 mg H3BO3 6 mg CuCl2 * 2 H2O 2 mg Adjust pH to 6.0 before adding water to 1000 ml total volume. Usage: 1 ml per liter medium. Multivitamin solution VL-7 (will be provided) Distilled water 180 ml Biotin solution (dissolve 10 mg biotin in 100 ml warm water) 20 ml Nicotinic acid 20 mg Thiamine dichloride 10 mg p-aminobenzoic acid 10 mg Ca-D-pantothenate 5 mg Pyridoxamine dihydrochloride 50 mg Cyanocobalamine (vitamin B12) 10 mg Sterile filter solution into screw-top bottles. Store in cool, dark environment! Usage: 1 ml per liter Medium. Dry mass determination In order to simplify and speed up the experiment, the specific bacterial dry mass is determined in a parallel experiment. Assuming that the dry weight per OD436 will not (significantly) change during the experiment, in the main experiment only the OD will be measured, and cells will be counted in the counting chamber. In parallel to the main experiment, two baffeld flasks, each containing 125 ml medium, are inoculated with 0.5 ml preculture each, and incubated with shaking at 37°C. At the end of the experiment, the two (fully grown) cultures are united, and the OD and the volume (250 ml cylinder) are measured (and taken note of). Afterwards, the dry mass is determined. • Thoroughly clean all weighing dishes (do not touch with your fingers but use tweezers to handle the dishes during cleaning), place in clean Petri dish, dry at 80 °C. So as to assure accuracy of your measurements, always weigh an empty container as a control.


• Precisely measure and note down the volume and OD of the culture to be examined (the more the better). Transfer cells into centrifuge beakers, counter-balance with water (including lids) (to precisely 0.5 g) and centrifuge at 12,000 rotations per minute to obtain a solid pellet. • Prepare 200 ml ammonium-acetate buffer, circa 50 mM, pH circa 6.5 (if necessary, adjust with acetic acid). This buffer evaporates completely as NH3 and acetic acid when heated (compare also Experiment T). • Immediately after the centrifuge has stopped, carefully remove the supernatant, re-suspend the cells in the ammonium acetate buffer. If necessary, gather cells from several beakers in one, wash cells (= centrifuge again) • Precisely weigh the dry containers. Write down empty weight. • Immediately after the centrifuge has stopped, remove the supernatant. (!Carefully!) transfer the cells to the weighing dish in 0.5 ml dist. water using a Pasteur pipette.

• Dry at 80 °C until the weight remains constant (1-2 days). • The final weighing has to occur immediately after the samples are taken out of the hot drying cabinet, as cells absorb considerable amounts of moisture while cooling down. Therefore, preset the analytical scale accordingly. • Thoroughly clean all weighing dishes. Calculate the dry mass per OD and ml. Main experiment - Inoculation of the test cultures In order to obtain a short lag phase and to be able to complete the experiment in a timely fashion, the test cultures are inoculated at 8 a.m., with a preculture that has been raised on the medium to be used in the main experiment with or without oxygen. 0.5 ml are added to the aerobic preparation and 0.3 ml are added to the anaerobic preparation before the tube is filled up with medium. Sterile pipettes (2 ml) are used for inoculation, as well as for taking samples throughout the experiment. Place test cultures in the incubator immediately and do not let them cool down in between.

Taking samples Immediately after inoculation, and at least every 60 minutes thereafter, 1 ml samples are taken from the aerobic cultures and examined for OD and cell number (max. once every hour). From anaerobic samples only the OD is measured. Temperature is an important parameter for biological turnover rates and should therefore remain constant. In no case leave cultures standing at room temperature. During the exponential growth phase, decrease the intervals at which samples are taken to 30 min. In order know when exponential growth is reached, the OD data have to be plotted semilogarithmically immediately (and compiled into a table)! Measuring OD The same photometer is used throughout the experiment to measure the OD. The aerobic cultures are examined using the one-cuvette-method (preferably glass cuvettes). To do this, set the photometer to zero using water as a blank. The sample is now measured using the same cuvette, after first drying the cuvette with filter paper. The OD should not exceed 0.3, otherwise the sample has to be diluted accordingly with 70 mM phosphate buffer (why not use water?). The screw-top tubes containing the anaerobically growing cells must not be opened until the end of the experiment (regardless of whether the OD exceeds 0.3!). These are instead measured against a tube filled with water. When the tubes are measured for the first time, the minimum absorption for each tube is found by turning the tube around in the photometer. This position is marked on


the tube with a felt pen, and the same position is used (after adjusting) to measure OD throughout the experiment.

Cell count The cell number is counted using a counting chamber (40x objective, phase contrast). For each count, at least 500 cells have to be counted, as statistical error at around √√√√N (factor for calculating the cell number per ml compare Schlegel). To count dilute in such a way that approximately 10 cells remain in each small square. End of experiment When the OD has remained static for 1 hour, the experiment can be stopped. The pH of every culture is to be measured and recorded at the end; sterile conditions are now no longer necessary. It is important to measure the final OD of the anaerobic cultures in a cuvette, after appropriate dilution, in order to compare the data. Evaluation Illustrate the measured growth parameters in both linear and semilogarithmic plots. Describe with words the growth in the oxic and anoxic assays. Calculate the maximum growth rates and the doubling time (formulas required!), the cell number, as well as the dry mass per ml and OD. Does the cell number per OD remain constant throughout the experiment? What is a first-order process? Write balanced chemical equations for the conversion of glucose with and without oxygen, as well as (simplified) for the synthesis of biomass. Which part of the glucose has been dissimilated, how much has been assimilated? What is the specific yield with and without oxygen? How do the results compare with your expectations? What is the dry mass of a cell? Assuming that the bacterium tested has the same mechanisms of energy conservation as E.coli, how much ATP is conserved with and without oxygen? What is the YATP? How many ATP-hydrolysis-cycles are necessary for the doubling of a cell? How much O2 is dissolved in an air-saturated medium (initial situation of anaerobic test culture), which proportion of the glucose could be converted with this O2?


Experiment R = Aerobic respiration Objective In this experiment the aerobic respiration by bacteria is studied. The O2 uptake of a suspension of washed cells in buffer is measured by means of an O2 electrode. Specific respiration rates (endogenous respiration and respiration with added substrates) are measured, and the affinity for O2 should be determined. Furthermore, the effect of an uncoupler is meassured.

Schedule Day 1 · Preparation of the medium · Inoculation of the bacterial culture · Preparation of all necessary solutions (cooling!) · Preparation of all devices · Familiarise yourself with the O2 electrode, data recording and calibration Day 2 · Washing and harvesting of the cells, cell pellets are suspended in buffer solution · Experiments with the O2 electrode

Materials Bacterial strain: Stock culture of strain M1 (or other)

For starting cultures: · 2 cylinders, 2 Erlenmeyer flasks (each 500 ml), magnetic stirrer · Complex medium for the strain to be studied · pH meter, calibration buffer, 1 M HCl · 2 baffled flasks (each 500 ml), felt cloth, aluminum foil for covering · Autoclave (coordinate use by the different groups!) · Bunsen burner, inoculation loop · Shaking incubator (set to an appropriate temperature, 30 or 37 °C)

For harvesting the cells: · Photometer: Shimadzu UV-1202, cuvettes (OS) · Refrigerated centrifuge with rotor for 250 ml cups (coordinate the use by the different groups!) · 2 centrifuge cups (250 ml) with cap · K-phosphate buffer, 50 mM, pH 7,0 (e.g. dilute from 1 M buffer) · Unsterile pipettetes (10 ml) · Robust test tube with rubber septum for the storage of the cells · Polystyrene containers with ice · Microliter syringe (100 µl) · Phase contrast microscope

For respiration experiments: · 2 O2 electrodes chambers (Bachofer/Hansa-Tech, incl. accessories) · N2 supply with needle valve


· Microliter syringes (25, 50, 100 µl) · Solutions: · KCl, 3 M · K-phosphate buffer, 50 mM, 500 ml (1 M buffer 1:20 dilution) · Complex-medium concentrate (50 x): 2 g Trypton + 1 g yeast extract in 20 ml H2O · Glucose, 1 M, 10 ml · Sodium acetate, 1 M, 10 ml · TCS (= Tetrachlorosalicylanilide, a decoupling agent or protonophor), 10 mM in methanol (poisonous, will be provided, handle with gloves!). Preparations with TCS are collected separately and are disposed in the central container

Preparation of the culture Per group 100 ml medium are inoculated in baffled flasks (500 ml). The cultures are shaken over night at optimum temperature.

Cell harvest For harvesting of the cells it is no longer necessary to work aseptically. First, the turbidity (OD436 against water) is measured, to estimate the growth yield. The photometer is set to zero with Buffer in a 3 ml-cuvette (if possible from glass) and the OD of the bacteria suspension is measured. At values higher than 0.3 the culture is diluted prior to the measurement. The suspension (approx. 100 ml) is transferred into a centrifuge beaker, which is counterbalanced against another cup filled with water on 0.5 g exact (both cups with caps!). The cells are centrifuged for 10 min at 4 °C and 8000 rpm (use the centrifuges only after proper instructions!). Afterwards the supernatant is decanted and rejected, the cell pellet is resuspended with buffer and centrifuged again (i.e. washed). The washed cells are then resuspended in 2 ml buffer, diluted to an OD436 of 40 (for OD measurement dilute 1:200) and transferred into a test tube, which is closed with a rubber septum and stored on ice. The conversion factors for dry weight (DW) and cell number per ml from the OD is taken from experiment G (provisionally you can assume 0.16 mg DW per ml for an OD = 1).

Measurement of the respiration rate Calibration and handling of the O2 electrode Do not damage the membrane on the bottom of the reaction chamber with the needles or Pasteur pipettes. The electrode chamber is filled with 3 M KCl over night. First of all the signals of the electrode in air-saturated and in N2-saturated buffer are recorded. For this, the electrode chamber is filled with air-saturated buffer and after temperature adaption (~ 3 min) it is flushed several times with fine air bubbles (take out some buffer with the Pasteur pipette and re-inject it from the gas phase). Wait until the signal is stable and write it down (100 % air saturation). Then flush fine N2 bubbles through the liquid, with a rubber septum loose on top (in order to reduce air entrance) wait for a stable value (max. 10 min) and write it down (0 % air saturation). After N2 saturation there is still O2 in the solution; complete O2 consumption by cells normally leads to smaller values! The calibration signals should be constant in each case and should lie in a "reasonable" range according to the tutor; if that is not the case a new diaphragm must be mounted. Determine the reaction volume of your system closed with the stopper by weighing the volume of the content after finishing an experiment.


For the calculation of the molar conversions you can assume that in air-saturated buffer at 37 °C 230 nmol O2 are dissolved per ml (240 nmol at 30 °C). Attach the signal with O2 saturation on the upper side of the display and try to write down the value before every measurement.

Endogenous respiration Air-saturated buffer is given into the reaction chamber which is then closed. Wait for a constant signal. Switch off the stirrer and add cells with a microliter syringe very carefully, so that the final OD is 1.0. The endogenous respiration rate should be small now or become so after a short time. The endogenous respiration rate is determined before each experiment and is substracted from the calculations of the substrate-specific rates.

Respiration of different substrates After determination of the endogenous respiration rates, test substrates are added, i.e. glucose, acetate (10 mM, each) or complex-medium concentrate (dilute the 50x medium concentrate 1:50). Record the oxygen uptake until complete consumption in order to be able to estimate the affinity for O2. Each rate should be measured twice. Effects of a decoupling agent With the substrate giving the highest respiration rates, test the influence of an uncoupler: Immediately after observing a constant rate, add 20 µM of TCS. Use a separate 25 µl syringe for this, afterwards rinse the reaction chamber several times! Solutions with TCS are collected in a separate container! Assessment Please show the following topics in your evalution: · Complete example of a respiration rate calculation from the measured curves (with a marked outprint of the results) · Illustrate your measured data in a table (respiration rates in nmol O2 min-1 mg-1 DW: endogenous respriation rates and respiration rates with the substrates and/or decoupling agent). · Most important: Summarise your results in your own words · Combine the rates with the data from the growth experiment to calculate the conversion rate of a cell. Which respiration rate did the cells have in your growth experiment while growing at maximum growth rate? Which respiration rate did the cells with complex-medium-concentrate have? · Please answer the following questions:

− How high is your own respiration rate if you take up food with 2500 kcal a day? − How does an O2 electrode work? − What is the KM value? − How do you estimate the affinity of the cells for O2? − How and where does TCS affect the respiration? To which extent was the respiration

"coupled" in the analysed cell suspension? − What are possible substrates for an endogenous respiration?


Experiment P = Phages Introduction Bacteria infecting viruses, called bacteriophages, are the most abundant biological entity on our planet with an estimated global number of about 1031. The phage abundance is direcly linked to the abundance of their hosts. They serve as controlling factors for microbial communities in the ocean,. According to the “killing the winner” hypothesis, phages prohibit “explosions” of single bacterial populations and keep microbial communities in balance. Furthermore, phages interfere with the microbial food web as they keep nutrients and carbon at a lower trophic level. The lysis of host cells causes the release of nutrients which directly fills up the pool of dissolved organic matter (DOM). Thus, the so called “viral shunt” has a wide impact on geochemical cycles.

In general, bacteriophages can exibit two lifemodes. During the lysogenic cycle, they integrate their genetic information into the host genome and reproduce as “prophages” together with the host cells. Due to environmental changes, their lifemode can become infective. This cycle is called lytic where cells are infected, phages are assembled within the cells and lyse the host afterwards. The released phages can subsequently infect new cells.

During the practical course, a phage induction experiment will be performed to demonstrate the presence of prophages in an infected bacterial culture. In a second experiment, a plaque assay will be set up to visualize the infection of host cells. Experimental schedule

Day Morning Afternoon 1st Induction experiment Preparation of material for plaque-assays 2nd Plaque-assays VLP counting via EFM 3rd Examine Plaque-assay results

1. Phage induction experiment Aim and general procedure During the induction experiment, the change from lysogeny to the lytic cycle will be demonstrated. The prophage inducing agent mitomycin C is used to catalyze an induction event. This antibiotic causes a DNA damage of the host and triggers the phage assembly. The development will be tracked by observing growth curves of bacterial cell cultures with and without induction. The released phage-like particles (VLPs) will be counted by epifluorescence microscopy (EFM). Material

- All material will be prepared in advance and distributed - 100 ml preculture of Paenibacillus glucanolyticus strain P073a (log-phase) - Mitomycin C (1.25 mg) - Fresh LB medium (sterile) - Two 50ml measuring cylinder (sterile)


Induction of prophages The culture will be incubated over night, shaken in the dark at 20°C to obtain a preculture in the early exponential growth phase. Next morning, the optical density will be measured at 580 nm (OD580) to get the starting point. After reaching a sufficiant OD580 , the culture will be split in two flasks. One part of the culture will be treated with mitomycin C (add 1.25 mg mitomycin C; final concentration 1 µg/ml) and the other one serves as a non-induced control.

After the treatment with mitomycin C for 30 min, the cells of the control and the treated culture will be washed twice by centrifugation (4000 rpm, 10 min, 22°C) and resuspended in 50 ml of fresh LB medium. Both subcultures will further be incubated at the given conditions and monitored hourly at OD580. Subsamples (1ml) will be taken before adding mitomycin C and at the end of the experiment after 8 to 10 hours to determine the viral abundance by counting. Therefore, each of the three samples will be filtered through 0.2µm filters and stored at 4°C until further use. Measuring the optical density The most accurate measurement range of the photometer is between an OD of 0.1 to 0.4. To match the range, the samples of the cultures have to be diluted before each measurement. The initial measurement is performed with a 1:2 dilution using fresh LB-Medium. After 2 and 3 hours a 1:5 dilution is used. Subsequently, the dilution facter is 1:10. Counting virus-like particles (VLPs) The counting of VLPs will be performed via epifluorescence microscopy. Briefly, the cell free phages from the subsamples will be filtered onto 0.02µm Anodisc filters and stained with the DNA staining dye SybrGreenI. Detailed instructions will be given during the practical course. Please answer the following questions in your protocols:

- Why do the induced free phage particles do not cause complete lysis of the culture? - Where do the viral-like particles in the non-induced control come from?

2. Plaque-assay Aim and general procedure Plaque-assays present a common technique for the detection, isolation and the enumeration of lytic bacteriophages. Within the practical course, plaque-assays will be performed with an E. coli strain that is sensitive for infection of the coliphage T4. Each group will get an phage lysate to determine the phage titer (phage/ml) via plaque-assays (plaque forming units, PFU, per ml). The plaque-assays will be performed using the agar overlay technique. Bacterial cells and phages are mixed with molten soft agar wherein host bacteria are then spread uniformly. This mixture will be poured on solid agar plates.


Material The required material for the plaque-assays will be prepared on the first day of the phage related experiments. They have to be prepared as follows:

- A culture of Escherichia coli (already prepared) - A phage lysate of coliphage T4 (already prepared) - 2 Flasks 250ml (sterile, see below) - 5 test glasses (sterile, see below) - 1.5ml reaction caps (sterile, see below) - NZCYM-Medium

Ingredient Weighted sample [g] Casein, hydrolysed 5 NaCl 2.5 Casamino acid 0.5 Yeast extract 2.5 MgSO4 x 7 H2O 1 Maltose 1 Add ddH2O to a total volume of 500 ml Adjust the pH to 7.5

The medium will be split in three flasks. The volumes should be 250 ml, 200 ml and 50 ml. For the following preparation of agar plates, 1.5% Agar (w/v) has to be added to 250 ml of the NZCYM-Medium. The soft agar is prepared by adding 0.65% Agar to 50 ml of the NZCYM-Medium. The rest of the medium will be used for liquid incubation of the E. coli preculture and serial dilutions of phage lysates. All prepared media, flasks and reaction caps have to be sterilized by autoclaving for 20 minutes at 121°C. Preculture of E. coli

To prepare the E. coli preculture, 100 ml NZCYM-Medium will be inoculated with 1 ml of an E. coli culture and incubated at 100 rpm over night at 37°C. This step will provide metabolically active cells for the following plaque-assay. Dilution series of phage lysates To avoid a complete clearance of the bacterial “lawn” on the agar plates, the phage lysate has to be diluted (see picture below). The phage lysate will be diluted with NZCYM-Medium. The dilution steps are done by adding 100µl of the lysate, respectively 100µl of the previous dilution, to 900 µl medium. The dilution series should vary from 10-1

to 10-10.


phage lysat 10-1 10-6 10-7 10-9

10-2 - 10-5

phage lysat 10-1 10-6 10-7 10-9

10-2 - 10-5

Preparation of plaque-assays The soft agar has to be boiled and tempered at 40°C in a water bath. After equilibration, five aliquots of soft agar (3 ml, each) will be prepared and kept in the water bath. A volume of 300 µl of the E. coli preculture and 50 µl of the different phage lysate dilutions (10-1, 10-7, 10-8, 10-9, 10-

10) are added to the soft agar, gently mixed and poured on the agar plates (see picture below). To disperse the top layer, the agar plates are carefully rotated. After solidifying, the plates will be incubated over night at 37°C. Finally, the plaques-forming units will be determined on the next day.

E. coli

Top-agar Phage T4

E. coli

Top-agar Phage T4

Please answer the following questions in your protocols:

- Why do we add maltose to the medium? - Why are there single colonies remaining on the agar plate after total lysis of the bacterial

“lawn” due to the use of undiluted phage suspensions?


E x p e r i m e n t T = Transport of salts across the cell membrane Aims In this experiment the uptake of various salts is analysed in order to draw conclusions about their transport mechanisms.

Approach In order to observe if solutes enter the cell, a high concentration of the compund of interest is added to the cell suspension. The increased osmolarity causes water export and shrinkage (plasmolysis) of the cells. Shrinkage is linked to an increase of the optical density, which can be observed and followed photometrically. When the solute enters the cell, it causes the cell to swell again, accompanied by a decrease in optical density. Time schedule Day 1 - Preparation of solutions and setting up of an overnight culture of strain M1 (or other bacterial strain) - Getting familiar with the needed equipment Day 2 Cell harvest and photometric experiments Required materials For inoculation of cultures: see Experiment R For the cell harvest: see Experiment R For photometric measurements: • Photometer: Shimadzu UV-1202 • Recorder or printer • Cuvettes made from optical glass, V = 4 ml, d = 1 cm • Graduated test tube, cylinder oder flask (10 ml) • Brandt-Transferpettor: 0.5 - 5 ml • Eppendorf-Comforpette: 100 - 1000 µl • Hamilton-syringe: 25 µl • Plastic spatula • Suspension of washed cells (OD = 10) • 50 mM potassium phosphate buffer, pH 7.0 5 M (or highest dissolvable concentration) solutions of the following salts in buffer: − Ammonium acetate − Sodium acetate − Potassium acetate − NaCl − KCl − Na2SO4 − K2SO4 • 10 mM tetrachloro-salicylanilide (TCS) will be provided TCS is toxic and should be handled with gloves only!


Inoculation of cultures: see Experiment R Cell harvest: see Experiment R After the last centrifugation, the cells are diluted in phosphate buffer in such a way that an optical density of 10 is obtained. Photometric measurements are performed at a wave length of 436 nm. Assay • 2.7 ml phosphate buffer • 0.3 ml cell suspension Mix assay and register the OD for a minute. Now add 0.13 ml of a salt solution, so that the final concentration in the cuvette will be 200 mmol/l. Mix as quickly as possible. The optical density is recorded until it does not change anymore. At the end: With salts that were not taken up (not decrease of initial turbidity increase with time), repeat the experiment and add TCS (20 µM) 1 minute after addition of the salt. For the report: With the help of your results, explain the transport mechanisms for ammonia, sodium, chloride, and acetate. Schematically illustrate the observed transport processes by drawings.

For the evaluation, please answer also the following questions:

- Which transport mechanisms of the cell do you know?

- Why can the membrane potential of the cell be relevant for the uptake of salts?

- How does an uncoupler work?

- What are the pK values of NH4+/ NH3 and CH3COOH/CH3COO-, and which compounds and

how much of them are formed when you dissolve ammonium acetate in neutral buffer?


Literature for student seminars: Mo (07. Dec): 1) Kane MD, Poulson LK, Stahl DA (1993) Monitoring the enrichment and isolation of sulfate-reducing bacteria by using oligonucleotide hydridization probes designed from environmentally derived 16S rRNA sequences. Appl. Environ. Microbiol. 59: 682-686 2) Lomans BP, Maas R, Luderer R, Op den Camp HJM, Pol A, Van der Drift C, Vogels GD (1999) Isolation and characterization of Methanomethylovorans hollandica gen. nov., sp. nov., isolated from freshwater sediment, a methylotrophic methanogen able to grow on dimethyl sulfide and methanthiol. Appl. Environ. Microbiol. 65: 3641-3650 Tu (08. Dec): 3) Klemps R, Cypionka H, Widdel F, Pfennig N (1985) Growth with hydrogen, and further physiological characteristics of Desulfotomaculum species. Arch. Microbiol. 143: 203-208. 4) Chen F, Wang K, Stewart J, andBelas R (2006) Induction of Multiple Prophages from a Marine Bacterium: a Genomic Approach. Appl. Environ. Microbiol. 72: 4995-5001 We (09. Dec): 5) Botero LM, Al-Niemi TS, McDermott TR (2000) Characterization of Two Inducible Phosphate Transport Systems in Rhizobium tropici. Appl Environ Microbiol. 2000 January; 66(1): 15–22. 6) Martínez-García S, Fernández E, Aranguren-Gassis M, and Eva Teira E (2009) In vivo electron transport system activity: a method to estimate respiration in natural marine microbial planktonic communities. Limnol. Oceanogr.: Methods 7, , 459–469

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