EXPERIMENT MANUAL FOR THE BIOCHEMICAL ENGINEERING LABORATORY
R. Mark Worden and Andrew B. Kinney Department of Chemical Engineering and Materials Science Michigan State University Revised August 2002 – Tim Bender
Table of Contents Laboratory Format______________ Format______________________ ________________ ________________ _______________ _______________ _______________ _______ 3 Report Format and Guidelines Guidelines _______________ _______________________ _______________ _______________ ________________ ____________ ____ 5 Laboratory Safety Regulations Regulations and and Laboratory Laboratory Policies ________ ________________ ________________ _____________ _____ 12 Aseptic Techniques Techniques for Culture Growth and Maintenance Maintenance ________________ _______________________ ___________ ____ 15 Batch Ethanol Ethanol Fermentation___________ Fermentation___________________ ________________ ________________ ________________ _______________ ________ _ 23 Enzyme Kinetics Kinetics _______________ _______________________ _______________ _______________ ________________ _______________ _______________ ________ 39 Plasmid Stability_______ Stability_______________ ________________ ________________ _______________ _______________ ________________ _______________ _______ 51 Immobilized Cell Biocatalysts Biocatalysts ________________ _______________________ _______________ ________________ ________________ ___________ ___ 64 Mass Transfer Transfer ________________ ________________________ _______________ _______________ ________________ ________________ _______________ _______ 84 Fermentation Fermentation Power Transfer_______ Transfer _______________ _______________ _______________ ________________ _______________ ____________ _____ 93 Membrane Filtration Filtration _________ _________________ ________________ _______________ _______________ ________________ ________________ ________ 100 Operating Procedures for Autoclave and Spectrophotometers _______________________ 119
Laboratory Format Lab Groups
During the first lab meeting, students will will be assigned into groups of three. Students will take turns being "group leader" for the main experiments, so that each student will serve as group leader for one experiment. Group structures structures of this type are common in industry. The purpose of having a group leader is to facilitate decision making and organization--not to shift the work load. All group members are expected to contribute equally on each experiment. Following the submission of each lab report, each student will be asked to turn in a confidential sheet describing his or her perception of the relative contributions of each of the group members. Lab Schedule
The laboratory component of CHE 481 will be divided into one demonstration experiment, which all students do simultaneously, simultaneously, and three experiments. Three periods will be allocated for each of the main experiments. The first period will be used for planning and preparation. During this period, students should learn how to operate the equipment, plan what runs they will do, and locate all of the supplies supplies they will need. It is especially important important to become familiar with the operation of the computer data acquisition acquisition systems during the first period. Although these systems have been designed to be relatively easy to operate, some learning and practice is required. Whenever possible, a trial run of the experiment experiment should be run during the planning period to identify any unforeseen difficulties. If additional supplies or special assistance (e.g., providing access to the laboratory outside ou tside of the standard times) is needed, the request should be made in writing to either the teaching assistant assistant or the instructor. instructor. The experiment will be carried out during the second and third periods. Results of the the experiment will then be written into into a short final report that will be due at the beginning of the next lab period (i.e., one week after the third lab period). Each experiment will be assigned assigned in advance of the first (planning) period. All group members are expected to have studied the appropriate chapter in the laboratory manual before the first lab period designated for that experiment. Unannounced quizzes may be given on this material. Part of the first lab period may also need to be used to clean up from the previous experiment. As examples, microbial cultures may need to be sterilized, and glassware may need to be washed and/or put away. Types of Experiments
There will be two types of main experiments: experiments: “existing” ones, which are already described in the laboratory manual, and “new” ones, which need further development before being included in the manual. During this term, term, some groups will be assigned a new experiment. Work done on new experiments will emphasize experimental planning and methods development, rather than acquisition and analysis analysis of large amounts of experimental data. Reports for new experiments should focus on describing the experimental methods tested and evaluating how well they worked. The suggested procedures should be written up in sufficient sufficient detail to enable someone unfamiliar with the equipment to conduct the experiment.
Procedures for existing experiments are described in the laboratory manual. A report format recommended for these experiments is given on the following pages.
Report Format and Guidelines Prereport Format
Experimental planning is critical if the experiment is to be carried out efficiently in the allotted time. To assist with the planning, a prereport will be written. The prereport must must be approved by the instructor or teaching assistant before the experimental work can begin. The prereport should be submitted to the Chemical Engineering Office (2527 EB) by 4:00 p.m. the day before the second lab period. The prereport format format is given below. Cover Page: This page should list the course number, title title and section number; the title of the experiment; the group letter, members, members, and leader; and the date submitted. submitted. There should be a blank for the instructor or teaching assistant to initial, verifying that the prereport has been approved, and a blank for the date of approval. Purpose of Experiment: The objectives of the experiment should be clearly stated. In other words, what will be learned by doing the experiment? Description and Operation of the Equipment: The experimental equipment should be described and illustrated. illustrated. The procedures for operating the the equipment should be outlined outlined in sufficient detail to show understanding. Theoretical Analysis and Sample Calculations: The pertinent chemical engineering principles and equations equa tions should be applied to the experimental system. Sample calculations should be shown using reasonable numerical numerical values. Include units in all calculations. calculations. Experimental Schedule and Flowchart: The dependent and independent variables should be identified, and the ranges of variables to be studied should be specified. The schedule of the experimental studies should be given, including the number of runs to be performed and the dates on which they will will be performed. The various steps involved in the experiment should should be summarized in the form of a flowchart that indicates which activities depend on others and the timing timing involved. For instance, to start taking data on a fermentation fermentation at 11:30 a.m., the reactor must be inoculated at 8:30 a.m., and to inoculate the reactor, the inoculum culture must be started late afternoon on the day before. Preparation of such a flowchart helps ensure that all the necessary steps get performed at the appropriate time. Supplies: All necessary reagents, glassware, and accessories should be listed, along with with the amount of each required. This list should should be developed early enough that the supplies may be located during the planning period. If something is not in stock, the instructor or teaching assistant should be notified. Sample Data Sheet: Where applicable, a sample of the actual data sheet to be used in the lab should be shown. Actual data sheets must be dated and initialed initialed by the instructor or teaching assistant at the end of each laboratory period. In several of the experiments, the data will be recorded automatically on an IBM-compatible IBM-compatible personal computer. Students
doing these experiments should provide a 3 1/2 inch disk formatted for high-density information storage. Safety Issues and Precautions: Precautions: Any potential hazards associated with the experiment should be described, along with the appropriate precautions. If microorganisms or hazardous materials are to be used, the proper methods of disposal should also be stated. Final Report Format
A short-report format will be used that focuses on the experimental results and discussion and leaves many of the details of apparatus, procedure, theoretical analysis, and calculations to the prereport. The short report is designed to concentrate co ncentrate the most important information in a brief document, and thus requires good organization and concise writing. The format of the short report is given below. Transmittal Memo: This memo, addressed to the instructor, should summarize the report report subject, the time interval covered, and list the authors. Title Page: This page should list the course number, title title and section number; the title of the experiment; the group letter, members, and leader; and the date submitted. Abstract: The abstract should should give a brief (100 to 150 word) overview of the the study, including the objective, the experimental approach, the key results, and the most important conclusions. There should be no graphs, tables, or references in the Abstract. Abstract. As it is often published separately from the rest of the report, the Abstract must be b e entirely self-contained (e.g., not contain references to the other parts of the report). Results: This section gives the important results of the study in a well-organized manner that is easy to follow. The text should clearly clearly tell what results are being presented and where in the report they may be found (e.g., the table or figure number). It should describe briefly how the results were calculated and refer to sample calculations in the appendices. Results should be presented in the format that most efficiently conveys the important information. Figures are usually preferable to tables tables for illustrating the effect of one or two independent variables, because they give a clear, visual picture of the trends. However, tables may be preferable when multiple variables must be considered simultaneously. simultaneously. All tables and figures need a number and a title (e.g., Figure 1. Effect of Impeller Reynolds Number on Power Number). Each table or figure should follow its first citation in the text. Experimental values should be compared to theoretical or literature values whenever possible. The following suggestions are offered for tables and figures: Tables All column headings should include units. un its. Include precision estimates if known (e.g., value ± standard deviation). In general, printouts of spreadsheets do not make satisfactory tables; word processors have excellent table-making capabilities.
Figures Axes should be labeled, including units. The independent variable should be on abscissa (horizontal axis). Include precision estimates whenever possible with error bars. Use symbols to mark data points and distinguish different curves. Maintain at least 1 inch margins. Use a straightedge or French curve c urve for neatness. Avoid unrealistic lines or curves through data points. If possible, use computer graphics to add a professional appearance. In cases where the results are obtained from the slopes of a large number of similar graphs (e.g., enzymatic reaction rates, and k La values), it is not necessary to show all of the graphs. Instead, show one or two typical ones, and then analyze the trends trends in the results. Discussion: In this section, the results should be interpreted in terms of whether the objectives were accomplished. The agreement between the experimental findings findings and the theoretical or literature results should be discussed. discussed. "Theoretical" results are those that can be derived from rigorous modeling equations. eq uations. "Literature" results are those that have been experimentally measured by others. Library work is often required to find such results for comparison. A rationale should be given for any discrepancies discrepancies between the expected and actual results. The significance of experimental error should be evaluated quantitatively. quantitatively. Key conclusions of the study should be stated and justified based on the results. Conclusions and Recommendations: The most important conclusions of the study should be reiterated in this section. Each of these conclusions should have already been stated and justified in the Discussion section. Recommendations for improving the experimental procedure, theoretical analysis, etc. should also be made in this section. Bibliography: The source of any quoted material or information that is not well known should be referenced. Use a standard format, such as Authors, Title of Paper or Report, Source, Date, Page Numbers. Any technical journal or text book will have examples of acceptable formats. Appendices: Appendices contain items of lesser importance such as sample calculations, tables of intermediate results, derivations, the prereport, and original data sheets (initialed by the instructor or teaching assistant). Each main item should have a separate Appendix designation and title (e.g., Appendix A: Prereport). An example of each type of calculation done for the report should be shown. Such calculations may be hand-written but should include typical values of the variables and units. Importance of Report Writing
One of the most important job job functions of practicing engineers is report writing. Consequently, writing effectiveness is one of the major criteria on which an engineer's performance is evaluated. Industrial representatives report that graduating engineers frequently have inadequate writing skills. This trend is probably due to the curriculum’s traditional emphasis on technical
training. In this class, one third of the the report grade will be based on quality of presentation, presentation, and two thirds on technical content.
Report-Writing Mechanics
The main objective of report writing is to efficiently convey to the reader the most important results of the experimental research. Too often, the results are contained in the report, report, but the report is not written so that the the reader can easily find them. Some of the keys to effective report writing are discussed below. Logical organization is essential essential for a good report. The report format given above provides a basic framework. However, each section must also be b e well organized internally. Developing a detailed outline before before writing the report can help in this regard. Such outlines are best developed in stages. In each sequential stage, additional levels of detail are added, dividing sections into smaller sub-sections. sub-sections. Once the outline is sufficiently sufficiently detailed, it is a simple simple matter to convert the individual ideas into sentences. The result is is a well-organized, well-organized, easy-to-follow report. When several people are working on different sections of the report simultaneously, redundancy and inconsistency in style often results. It has been said that the quality of a report varies inversely with the number of writers writers and editors. For this reason, it is often often useful to designate one person to do the final editing of the report. This person is responsible for integrating the various components into a cohesive final product. Writing style is also important. important. A "Fog Index", shown below, has been developed by Robert Gunning (1) to estimate how difficult a passage of writing is to follow. F = 0.4 (W/S + 100 P/W) where
F = Fog Index: the number of years of schooling required for a reader to understand what has been written. W = the number of words in the passage being evaluated (W should be at least 100). S = the number of sentences in the passage being evaluated. P = the number of polysyllabic words (three or more syllables) in the passage. Words having three syllables because a suffix was added (e.g., "fastening") are not counted.
Popular magazines have an F value of about 8 to 12. While this range may may be a few points too low for a technical report, passages with an F value greater than 17 often cannot be understood by a general reader. The Fog Index points out that short sentences and simple words help the reader follow what is being read. The unnecessary use of long, convoluted sentences and big words does not impress readers--it only hinders their understanding! In proofreading, then, one should eliminate words that add nothing to the meaning of the sentence and "pretentious" words that could be replaced by simpler words.
Professional engineering reports are expected to be typed and have correct grammar and spelling. These standards will be required in this this course. However, because equations and tables are difficult to type, they may be written neatly in pen. Excellent word-processing software is supported by the College of Engineering. Most word processors processors now have spelling checkers, and some check grammar as well. However, there is no substitute for detailed proofreading. proofreading. Each group member should proofread the report before it is submitted. Computer-based statistical analysis and graphics also can add professionalism with minimal effort. For example, regression coefficients for linear linear fits and standard standard deviations of replicate data can be calculated automatically by most spreadsheet packages.
Bibliography
1. Gunning, R. How to Take the Fog Out of Writing, Dartnell Corp., Chicago, IL., 1964.
CHE 481 Laboratory Report Grade Sheet Group Letter ______
Group Leader ________________________________
Prereport (25)
Technical Content (17) Objectives, Equipment Operation and Safety (4)
______
Theoretical Analysis (4)
______
Experimental Plan, Schedule (9)
______
Presentation Quality (8)
______
Final Report (75)
Technical Content (50) Abstract (10)
______
Quality, Quantity, and Presentation of Results (15)
______
Discussion of Results and Error Analysis (15)
______
Conclusions and Recommendations (10)
______
Presentation Quality (25)
Total Grade (100)
______
_____
Oral Presentation Rating Form
Student’s Name _______________ ___________________________ ____________ Course_________________ Course_________________ Date _____________ _____________ Presentation (75 points) Organization (25 pts) • Introduction- oriented the audience to help them understand the topic and problems • Main ideas (well-defined, distinct, well-supported) • Transitions (signal movements and/or phrases, logical order, smooth flow) • Summary/Conclusions (main message(s) clear and distinct) • Overall impression
_____ _____ _____ _____ _____
Delivery (35 pts) • Appropriateness (language adapted to audience, not too simple, not too complex) • Clarity (language used to promote understanding) • Stance, movements, gestures • Voice quality, pitch, volume, speaking rate • Eye contact (constant, occasional or rare) • Control (situation under control by speaker) • Overall impression
_____ _____ _____ _____ _____ _____ _____
Visual Aids (15 pts) • Size/clarity (easily viewed, not too much detail, not too few or too many, assist understanding of presentation) Integration/subservience into speech/smooth transitions from slide to slide • Integration/subservience • Overall impression
_____ _____
Participation (5 pts) • Distribution of speaking among group members
_____
Technical Content (20 pts) • Scope of experiments (adequate number and type of experiments) • Data analysis (careful, thoughtful consideration of data and their meaning) • Error analysis (sources of error, statistical treatment of data) • Relationship between theory & experiments (theory described & used to interpret data)
_____ _____ _____ _____
TOTAL SCORE
_____
_____
Instructor Comments/Suggestions
_______________ _______________________ ________________ _______________ _______________ ________________ _______________ _______________ _______________ _________ __
Laboratory Safety Regulations and Laboratory Policies Safety in the laboratory is the highest priority. Each student is responsible for for understanding and following the safety regulations described below. Questions regarding the regulations should be referred to the instructor. GENERAL LABORATORY SAFETY
1.
Know the location of and how to use use of the following following safety safety equipment: a. b. c. d. e. f. g. h. i.
safety shower fire extinguishers fire blanket first aid kit eye wash station gloves chemical spill kit Materials Safety Data Sheets Antiseptic solution
2.
Immediately report all hazardous situations, situations, accidents or injuries to the the instructor. instructor.
3.
Wear protective glasses or goggles with side-splash protection in the laboratory at all times.
4.
Refrain from from activities activities that that might might accidentally introduce hazardous material into your mouth or eyes in the laboratory (e.g., eating, drinking, smoking, applying cosmetics).
5.
Learn and avoid and cope with the hazards specific to the equipment before you use it.
6.
Learn the hazardous characteristics characteristics of the materials materials involved in the experiment experiment before you use them, and take proper precautions during their their use. The Material Safety Data Sheets (MSDS) located in a notebook in the laboratory have this information and should be consulted before the experiment is begun.
7.
Use and store all materials materials in a location appropriate for their their hazards.
8.
Maintain a clean and organized work area at all times.
9.
Become safety-conscious, safety-conscious, and assist others in maintaining a safe safe laboratory environment.
10. Avoid open-toed shoes when working in the laboratory. Lab coats are recommended. 11. Label all containers with the contents, contents, date, group letter, and initials initials of person preparing.
BIOHAZARDS
The Biochemical Engineering Laboratory uses only microorganisms that are non-pathogenic (i.e., do not cause sickness). As a result, the lowest lowest level of precaution, Biosafety Level 1 (BL1), is sufficient. The BL-1 guidelines, also known as Standard Microbiological Practices, are listed below. 1.
Work surfaces surfaces are decontaminated once a day and after any spill spill of living cells.
2.
All materials materials contaminated with with living cells (e.g., petri dishes and disposable pipettes) pipettes) are decontaminated (sterilized) before disposal. A central location will be designated for materials that need to be sterilized before disposal.
3.
Do not mouth pipette. Use a mechanical device to draw and dispense liquids with a pipette.
4.
Do not eat, drink, smoke, apply cosmetics, etc. in the laboratory. Keep food out of the laboratory.
5.
Persons should should wash their hands after handling materials that might might be contaminated with with living cells and before they leave the laboratory.
6.
Try to minimize the formation formation of aerosols that might carry living living cells. cells.
7.
Laboratory coats are recommended recommended to prevent contamination contamination or soiling of street street clothes. clothes.
Laboratory Policies and Procedures Decontamination of Equipment: Materials coming in contact with with live cultures should be sterilized after use. Glassware, Petri dishes, dishes, disposable pipettes, pipettes, syringes, etc. will be sterilized by autoclaving. The middle shelf of the autoclave rack (located next to the autoclave in 394 EB) is used to collect materials needing autoclaving. Tabletops and reusable materials that cannot withstand autoclaving (e.g., the plastic reservoirs for the membrane filtration experiment) will be sterilized using an antiseptic solution. Biohazard Bags: Prior to autoclaving, small, non-sharp items (e.g., disposable plastic pipettes, pipette tips, and filters) may be placed in the orange bioh azard bag located in the culture-transfer area on the south wall of 388 EB. After autoclaving, the bags should be taped closed, wrapped in a black plastic bag, and disposed of in the trash can. Sharps Container: Waste materials with sharp edges or points should be placed in the sharps container located in the culture-transfer culture-transfer area. Disposable syringes must also be discarded in the sharps container even if they do not have a needle attached. attached. Broken glass that is not contaminated with microorganisms may be discarded in a specially designated bucket located by the door of 388 EB. Hazardous Wastes: Hazardous wastes should be discarded into the specially designated plastic waste barrel located in the walk-in hood of 394 EB. The contents and amount of the waste should be logged onto the attached sheet. Before emptying the waste into the barrel, please check for incompatibilities with other wastes in the barrel. If in doubt, please ask the teaching assistant or instructor. Glassware Cleaning: After use, all glassware should be cleaned and returned to the appropriate storage storage cabinet at the the earliest earliest opportunity. opportunity. Suggestions for for glassware glassware cleaning are posted by the RO water tap on the south wall of 388 EB. After the final RO water rinse, the glassware should be allowed to dry in the drying racks located in the walk-in hood of 394 EB. Desktop and Bench Space: The desks, tables and lab benches need to be left clear for use by students. Please put coats on the coat racks and backpack s on the floor out of the way. The lab bench in the center of 394 EB has been designated for medium preparation. Other types of lab procedures should be performed elsewhere.
Aseptic Techniques for Culture Growth and Maintenance Introduction Sterility is a critical critical requirement for many industrial bioprocesses. Unwanted microorganisms (i.e., contaminants) can reduce the yield of a desired product by competing for nutrients or metabolizing the product itself. They may also introduce some undesirable compound or property into the product. Aseptic (i.e., sterile) operation is achieved by first sterilizing the equipment and its internal contents and then preventing contamination by unwanted microorganisms. The objective of this demonstration experiment is to learn aseptic techniques for culture transfer. Sterile liquid and solid growth growth media will be prepared and then inoculated (i.e., seeded) with a pure culture of Escherichia Escherichia coli. coli. After incubation in a constanttemperature incubator, the size, shape and color of the resulting cell colonies will be inspected to identify the presence of contaminants.
Background Sterilization is most commonly accomplished in industrial practice by hea ting with saturated steam. The Biochemical Engineering Laboratory has a computer-controlled computer-controlled steam autoclave that will maintain a desired internal temperature for a specified length of time. For small volumes of material, material, sterilization for 15 minutes at 121°C (15 psig) is generally sufficient. Longer heating times are needed for large objects or if sporeforming organisms are known to be present. Spores are heat-resistant, dormant dormant cells that can survive harsh conditions. Microbial cultures are transferred by adding a small amount o f a pure culture of the desired microbe to sterile growth medium. Occasionally microbial contaminants may find their way into the growth medium and grow along with the desired species. If undetected, the contaminants may cause problems. In extreme cases, trace quantities of potent toxins produced by unwanted microbes may be life-threatening. Therefore, it is good practice both to attempt to avoid contamination and to occasionally verify that the culture is still pure. One method to initiate a pure culture is to begin with a single cell. If a very dilute suspension of cells is spread onto the surface of sterile agar in a petri dish, each cell will grow into a single, macroscopic colony that tha t is physically separated from other colonies. If cells from only one of these colonies are then used to initiate a new culture, that culture should be pure.
This plating technique can also be used to check for contamination. Different species often have different colony morphologies (i.e., (i.e., appearances). Thus, if colonies with different sizes, colors, shapes, edge characteristics, etc. develop on the plate, the culture was probably contaminated. However, this test has limitations. There may be contamination even when there is no n o obvious variation in the colony co lony characteristics, and some pure cultures can show variety in colony morphologies. Once a pure culture is established, it may be used for some bioconversion step, stored by freezing or lyophilization (freeze-drying) or simply maintained in a refrigerator. Refrigerated cultures have a finite finite lifetime that varies varies with the species. Consequently, the culture must occasionally be regrown. regrown. This periodic “culture transfer” procedure procedure may be performed in any of the following four ways: liquid-to-liquid transfer, liquid-to-solid liquid-to-solid transfer, solid-to-solid transfer and solid-to-liquid transfer.
Experimental Equipment and Procedure All growth media will be prepared by the students, working in groups. Each group should prepare seven sterile LB-agar plates and seven 125-mL Erlenmeyer flasks, each containing 50 mL of sterile sterile LB broth. Instructions for preparing preparing the plates and liquid media are given in Appendix 2. Then, following a demonstration of aseptic culturetransfer techniques by the instructor, each student should do the following transfers: 1. With a wire loop, aseptically streak E. E. coli cells taken from a single colony on the agar plate provided by the instructor onto a sterile agar plate. 2. With a wire loop, aseptically transfer E. E. coli cells from a single colony on the agar plate provided by the instructor into sterile LB broth. 3. With a wire loop, aseptically streak E. E. coli cells taken from the liquid culture provided by the instructor onto a sterile LB plate. 4. With a 10 mL, sterile, disposable disposable pipet, aseptically transfer transfer E. E. coli cells from liquid culture into sterile LB broth. All culture transfer operations will be done in the culture transfer area in the SW corner of 3262 EB. Because it is partially enclosed, this area has less air currents that could carry microbes into the flasks or plates. To maximize efficiency, all students will do one type of transfer before going on to the next. The instructor will supply supply the pure cultures. After inoculation, the flasks and agar plates should be labeled with the contents, date, group, and person doing the transfer. All plates and flasks will be placed in the large incubator located in the NE corner of 3269 EB and grown at 35°C. Agar plates plates should be always be stored upside down to prevent condensation from dripping onto the agar surface. Arrangements should be made with the teaching assistant to transfer transfer flasks and plates to the refrigerator at the proper time. This strain of E. E. coli grows up overnight at 35°C. However, the plates should be allowed to mature for a couple of days to allow slower growing contaminants time to develop. During the second laboratory period, the plates should be examined for evidence of contamination. Individual colonies should be readily apparent on part of the plate. The
morphologies of the colonies should all be identical. Significant differences in morphology between colonies is is suggestive of contamination by unwanted species. To check the purity of the liquid cultures, each student should streak a sample from one of the liquid cultures that student inoculated during the first period onto a sterile agar plate. These plates should be properly labelled and then incubated upside down at 35°C. Arrangements should be made with the teaching assistant to transfer the plates to the refrigerator at the proper proper time. At this point, students should begin preparation for the the next experiment. During the third laboratory period, period, the plates streaked during the the second week should be examined for evidence of contamination. All plates and flasks should be autoclaved following use. Flasks should be washed as described on a sheet by the wall sink in 3262 EB, allowed to dry in the drying rack, and finally returned to the storage storage cabinet. After autoclaving, the petri dishes should be wrapped in an orange biohazard bag, then a black plastic bag, and finally discarded in the trash can.
Appendix 1: Equipment and Reagents Needed: Needed: E. coli culture on LB plates E. coli culture in LB broth Seven 125 mL Erlenmeyer flasks Seven sterile, disposable Petri dishes granular agar LB broth components (alternatively, pre-mixed LB broth powder) tryptone yeast extract NaCl Bunsen burner wire transfer loop sterile pipettes pH meter NaOH solution 95% ethanol
Appendix 2: Details of of Experimental Procedure I.
Preparation of sterile LB broth in an Erlenmeyer flask A. Weigh out enough pre-mixed LB broth powder to make 500 mL at a 20 g/L concentration. If the pre-mixed pre-mixed LB powder is not available, prepare the LB broth according to the following recipe: Tryptone..................................10 g/L Yeast Extract.............................5 g/L NaCl ........................................ 10 g/L If available, you may use the prepared LB broth reagent powder instead of the recipe above.
B.
Dissolve the reagents in the the proper amount of distilled water.
C.
If the pH is not within 0.2 pH units of 7.0, adjust the pH into this range with the pre-mixed NaOH solution. The pre-mixed LB-broth powder comes already pH adjusted.
D. Distribute 50 50 mL of the LB broth into each of seven 125 Erlenmeyer flasks. E.
II.
Insert a foam plug (or a wad of non-wetting cotton) into the mouth of each flask. The plug prevents air-borne microbes from entering the flask flask once it is sterilized. Enough of the plug must protrude protrude from the mouth of the flask (about ¾ to 1 inch) to allow e asy removal during culture transfer.
Preparation of LB-agar solution A. Place 150 mL of LB broth into a 250 mL Erlenmeyer flask. flask. B.
Add enough granular agar to the LB broth to give an agar concentration of 20 g/L. Note: the agar will not dissolve until the solution has been heated to boiling during the autoclaving cycle.
C.
Insert a foam plug (or (or a wad of non-wetting cotton) into the mouth of the flask. The plug prevents air-borne microbes from entering the flask once it is sterilized.
III. Autoclaving the LB broth and LB-agar solutions A. Autoclave the flasks containing LB broth and LB-agar solutions at 121°C for 15 minutes, as described in the last chapter in the laboratory manual. B.
After autoclaving, the solutions may be cooled by partially submerging the the flasks in cold water. However, the agar solution must not be cooled below 50°C, and must be swirled continuously during during cooling. Otherwise, the agar will solidify on the inner wall of the flask.
IV. Pouring the Agar Plates A. This process of cooling the hot agar to 50°C while swirling is called “tempering”. If very hot solution is poured into the plates, too much condensation will occur on the inside of the lid. Drops of condensation can cause problems (e.g., mixing of cells from different colonies). If the agar begins to solidify before you are ready to pour the plates (its freezing point is 42 °C), reheat in the microwave. Once the agar has solidified, it must be reheated to 100°C to be liquefied again. B.
Remove the foam plug from the the mouth of the flask and “flame” the mouth of the flask (i.e., rotate the lip of the flask through a Bunsen burner flame).
C.
Lift the lid of a plate, and pour enough agar solution into the plate to just cover the bottom (about 15-20 mL). Replace the lid. lid. Any bubbles that may have formed during pouring may be broken by gently swirling the plate.
D. Stack the next plate to to be poured on top of the one just poured to insulate the lid of the plate and hence minimize condensation inside the lid. E.
Repeat the previous previous two steps until all seven plates are poured. Place something something on top of the stack (e.g., paper) to insulate and minimize condensation in the top plate.
F.
Immediately after pouring the plates, rinse the remaining remaining agar down the drain with plenty of hot water. Once the agar has solidified, solidified, it can be difficult to remove from the flask.
V. Streaking a plate from liquid or solid culture A. Light the Bunsen burner in the culture-transfer area, and heat the wire loop until it glows red. B.
C.
Transferring cells from liquid culture: 1.
Remove the foam plug from the flask, flame the lip of the flask, and dip the loop into the liquid culture.
2.
Remove the wire loop. Flame the lip of the flask, and replace the foam plug. During this procedure, do not put the foam plug down, and handle it only by the top to minimize chances of contamination.
Transferring cells from solid culture: 1.
Raise the lid of the plate containing the culture, and cool the wire loop by touching it onto the agar where there is no growth.
2.
Gently slide the loop through a single colony to pick up cells. Transferring cells from only one colony minimizes the cha nces of contaminating the new culture culture medium. Avoid cutting into the agar with the wire loop.
3.
Replace the lid of the plate. Note: because cells are microscopic, it is not necessary to get enough cell mass on the wire loop to be seen by eye.
D. Streaking the plate (Refer to Figure 1) 1.
Raise the lid of the plate to be streaked, and lightly streak (i.e., wipe) the wire loop back and forth across the surface of the agar to spread the cells
across one area of the plate (e.g. region 1). Avoid any activity that could create air currents in culture-transfer area. 2.
Close the lid and heat the wire loop to glowing again. Lift the lid, cool the wire loop in the agar, and then, starting with the loop in region 1, make several parallel steaks that extend into a fresh region of the agar (region 2). This process spreads out (dilutes) a portion of the cells depo sited in region 1 over region 2.
3.
Repeat the previous step once or twice more, starting with the wire loop in a the region just streaked and moving moving into a new area. The objective is to sequentially dilute the cell concentration with each streaking. Eventually, the cells are sufficiently dilute that individual colonies, each derived from a single cell, can be observed. Then, one colony may be picked for inoculation purposes, and the morphology of different colonies may be compared to check for contamination.
4.
Replace the lid. Store the plates upside down to prevent any condensation that may occur from dripping down onto the agar.
5.
Leave the plates uncovered (upside down) overnight to allow any water that may seep out of the agar (a process called syneresis) to evaporate. Then, stretch a band of Parafilm around the edge of the plate to help keep out airborne microbes.
6.
Incubate at 37°C until colonies develop.
VI. Inoculating liquid medium medium in an Erlenmeyer Erlenmeyer flask from solid culture A. Find a clean, still area to minimize the chances of contamination from the air. B.
Light a Bunsen burner, and heat the wire loop until it glows red.
C.
Raise the lid of the plate containing the culture and cool the wire loop by dipping it into the agar where there is no growth. Then, remove part of a colony using the loop. Replace the lid.
D. Remove the foam plug from the flask containing the sterile sterile liquid medium, flame the lip of the flask, and an d dip the wire loop into the sterile medium. Remove the wire loop. Flame the lip of the flask, flask, and replace the foam plug. VII. Inoculating liquid medium in an Erlenmeyer flask from liquid culture A. Familiarize yourself with with the operation of the pipette bulb using non-sterile pipettes. You should be able to draw and release liquid using only the thumb and first two fingers of one hand.
B.
Find a clean, still area to minimize the chances of contamination from the air, and light a Bunsen burner.
C.
Open the plastic wrapper wrapper on the pipette enough to attach the pipette bulb onto sterile pipette.
D. Peel the wrapper off the pipette from top to bottom while holding the pipette bulb with the other hand. Do not allow any part of the pipette that will enter the flask to touch a surface (or hands) that could be contaminated with microbes. E.
With the other hand, pick up the flask containing the liquid culture. culture.
F.
Wrap the little finger of your pipette hand around the top of the foam plug (without touching the lip of the flask with your finger) and gently remove the plug. Continue holding the plug with your little finger.
G. Flame the lip of the flask, and insert the pipette into the flask. H. Draw into the pipette only the amount of culture needed for the inoculation . A typical volume is between 1 and 5% of the sterile-medium volume. To minimize the possibility of contamination, avoid touching the neck of the flask with the pipette. I.
Remove the pipette, flame the lip of the flask, and replace the foam foam plug.
J.
Repeat steps steps F and G for the flask containing the sterile medium, and then release the liquid culture into the sterile medium. To minimize the possibility of contamination, avoid touching the neck of the flask with the pipette.
K. Remove the pipette, flame the lip of the flask, and replace the foam plug. L.
Place the wet end of the pipette into the biohazard bag before removing the the suction bulb (Removing the bulb can cause unintentional drips). Drop the pipette into bag.
M. Sanitize any spills or drips with antiseptic antiseptic solution.
Figure 1. Schematic Diagram of a Typical Plate Streaking Streaking Pattern.
Batch Ethanol Fermentation Introduction Fermentations may be thought of as chemical reactions catalyzed by living cells. cells. A variety of products can be produced by fermentation, such as pharmaceuticals, organic acids, and alcohols. However, to be commercially viable, the bioprocesses must be economically competitive with alternative processes, such as petrochemical manufacturing. Advances in recombinant-DNA technology allow fermentations to mass produce chiral and complex biomolecules (e.g., such as human insulin) more economically than by other means. The most widely known fermentation product is ethanol. Batch yeast fermentations have been used for hundreds of years to produce alcoholic beverages. (Ethanol can be produced from a variety of plant-derived raw materials, including agricultural wastes. There is currently a small market for ethanol as a fuel additive, but for fermentation ethanol to successfully compete with petroleum, additional improvements in the process economics are needed. The use of anaerobic bacteria, such as Zymomonas mobilis, mobilis, that grow faster and give higher ethanol yields than the conventionally used yeasts yea sts could provide such an improvement. (This experiment previously used Z. used Z. mobilis, mobilis, but in 2001 we switched to Saccharomyces cerevisiae (brewers’ yeast). The reference to Z. to Z. mobilis has been left in for its informational value.) The objectives of this experiment experiment are to carry out a batch ethanol fermentation using S. cerevisiae and to study the kinetics and stoichiometry of this fermentation.
Background Ethanol has several attractive features as an alternative fuel. As a liquid, it is easily transported. It has a heating value 2/3 that of gasoline, and it can be blended with with gasoline to increase the octane rating of the fuel. Ethanol was used as a primary primary fuel before and during World War II but was later replaced by cheaper petrochemical products. The huge fluctuations in the price of petroleum within the past twenty years have made commercial production of fermentation ethanol a more attractive, but still risky, venture. The perceived need for the US to have a stable and renewable energy base may spur an increase in the production of fuel ethanol in coming years. Innovative processing strategies are required to make ethano l production economically competitive with petrochemical petrochemical products. Recent developments, such as continuous fermentations with cell recycle and vacuum operation, have provided a twelve-fold increase in productivity over conventional processes (Bailey (Bailey and Ollis, 1986). Tower fermenters are used to retain flocculent microbes, eliminating the need for auxiliary separation devices (Bailey and Ollis, 1986).
Fermentations are usually carried out using a single (pure) microbial culture to assure a high-quality product. Contamination of a fermentation by unwanted microorganisms microorganisms is avoided by initially sterilizing sterilizing the fermenter and reactants. Then, precautions are taken to prevent entry of unwanted microbes. To initiate the fermentation, the sterile fermenter is inoculated by aseptically adding a small quantity of the desired species. After inoculation, a batch fermentation fermentation goes through four phases: the lag phase, the exponential growth phase, the the stationary phase, and the death phase. During the lag phase, the cells adapt to their new environment; little growth occurs during this phase. In the exponential growth phase, the cells grow rapidly, dividing with a constant doubling time. As a result, the cell concentration increases increases exponentially. This phase continues until changing conditions in the reactor reactor affect the growth rate of the cells. As examples, substrate concentrations may drop to rate-limiting levels, or product concen tration may increase to inhibitory levels. Cell growth eventually ceases in the the stationary phase. During this phase, cells often consume stored ene rgy reserves to sustain their viability. Finally, in the death phase, the cells die, typically at an exponential rate. A fermentation is monitored by measuring the substrate, product, and cell concentrations during the fermentation. Colorimetric methods are are frequently used, whereby samples are combined with enzymes to catalyze reactions that form colored products. The optical density (OD), also referred to as absorbance, of the colored product is then measured and a nd compared to a calibration curve constructed using known standards. Cell concentrations are usually measured by either the OD of the fermentation broth or the cell dry weight. The reduction in intensity of light passing through a cell suspension is primarily due to diffraction of the light light by the individual cells. These turbidity measurements are quicker and sensitive at lower concentrations than dry-weight determinations, but they give relative concentration da ta that must be converted to true cell concentration units using a calibration curve. Dry-weight assays are imprecise imprecise at low concentrations, but they are useful late in the fermentation when the cell concentration is highest. Samples taken at this time time may be used to develop a calibration curve for the turbidity assay, and simultaneous measurements of the glucose, ethanol and cell concentration at the end of the fermentation may be used to calculate the overall carbon and electron balances. The dry-weight assay entails removing a known volume of fermentation broth, separating the cells from the nonvolatile salts and substrates in the medium, and then drying and weighing the cells. The concentration is calculated by dividing the dry cell weight by the initial volume of the sample.
Experimental Equipment and Procedure (Note: the following paragraph is again referring to labs prior to 2001. The discussion would be similar for the brewers’ yeast that you will be using.) (Zymomonas mobilis (ATCC 10988) is a rod shaped, anaerobic, gram negative bacterium that converts glucose to ethanol by the Entner-Doudoroff pathway. Z. mobilis has been suggested for commercial ethanol production because it has a higher growth rate and -1 ethanol yield than yeasts; its specific growth rate at 30°C is about 0.40 h (Worden, 1982). Z. mobilis is often used to brew alcoholic beverages in tropical climates.
The overall reaction catalyzed by the cells is shown below: cells ammonia + glucose → ethanol → ethanol + carbon dioxide
(1)
Z. mobilis grows well between pH values of 5.5 and 8 in solutions containing up to 20% glucose. The carbon dioxide produced by the fermentation acidifies the medium, eventually inhibiting the fermentation unless the pH is maintained by adding base or buffering (Worden, 1982). The New Brunswick BioFlo IICtm fermenter is capable of controlling the pH, oxy gen concentration, temperature, and agitation agitation rate. However, because this fermentation is anaerobic and will be buffered, only temperature and agitation control are needed. A suitable fermentation medium consists of 2 % (by weight) gluc ose, 2% peptone and 1 % yeast extract (a powdered nutrient mix obtained from yeasts) at a pH around 7. Enough medium should be prepared and sterilized to grow two inoculum cultures cu ltures (100 mL each) and run the fermentation in the stirred-tank bioreactor bioreactor (1 L). The glucose may be autoclaved separately from the peptone and yeast extract to prevent darkening of the solution. An apparatus suitable for autoclaving 500 mL of liquid and then aseptically aseptically transferring the liquid to another vessel is shown in Figure 1. To make 1 L of medium, the glucose can be autoclaved in 500 mL of water in one flask, and the peptone and yeast extract can be autoclaved in 500 mL of water in another flask (after adjusting the pH to 7.0). The fermentation vessel may be autoclaved as shown in Figure 2. Although this figure shows a pH probe, automatic pH control is not essential, since the medium is buffered. The medium may be autoclaved a utoclaved outside of the fermenter and then aseptically transferred to the fermenter prior to inoculation. Two 250 mL Erlenmeyer flasks, each containing 100 mL of the same medium to be used in the fermenter, should also also be autoclaved. Experience has shown that it is not critical to autoclave the glucose separately from the other components for these “inoculum culture” flasks. The flasks should be capped with foam foam plugs to allow passage of steam during autoclaving while keeping out unwanted microbes. During the fermentation, samples (≅1 mL) should be taken periodically for glucose and ethanol assays. It is especially important important to take initial and final final samples in order to to do an overall electron balance on the fermentation. fermentation. The initial sample should should be taken just after the reactor has been inoculated, and the final sample should be taken the day after the fermentation is run. run. The liquid medium must be separated from from the cells immediately after sampling to stop stop the reaction. The microcentrifuge can be used to spin spin down the cells. The clarified liquid medium should then be transferred to a clean clean Eppendorf tube using a pipette, labelled, and stored stored in the refrigerator. Samples may also be drawn at a few other times times to track track the progress of the fermentation. However, it can be difficult to
accurately measure changes in the glucose concentration early in the fermentation, because the percent change is quite small. The glucose and ethanol assays are performed using commercially available, colorimetric assay kits. An aliquot of cell-free liquid liquid medium is mixed with with the appropriate reagent, and the reaction is allowed to proceed for a specified time. time. The OD is then measured at a specified wavelength. Detailed instructions for for the assays may be found found either in Appendix 1or in the instruction sheet for the assay kit. A calibration curve must be made up for the glucose and ethanol assays by assaying standards (samples whose concentration is accurately known). A calibration curve is made by plotting the OD values of the standards vs. their concentrations. It shows the correlation between concentration and OD, so that the concentrations of the fermentation samples (whose concentrations are not known) ca n be determined from their measured OD values. Sometimes standards are included with the assay kit. If not, they can be made up by accurately diluting a known amount of either glucose or ethanol to a known final volume using the analytical balance and volumetric pipettes and flasks. flasks. The concentrations of the standards should lie in the “linear range” of the assay, which is given in the assay instructions sheet. Fermentation samples will likely have to be diluted down into the linear range before doing the assays. If the OD values are outside this range, the results may be inaccurate. inaccurate. The standards should be assayed at the same time as the fermentation samples for greatest accuracy. The initial and final samples samples should be assayed at least in duplicate to improve accuracy and get a sense of the precision of the assay. The OD of the fermentation broth should be measured about every 20 minutes during the exponential growth phase to monitor the specific growth rate. Also, the cell concentration should be measured in duplicate (25.0 mL per assay) at the end of the fermentation (the day after the the fermentation is run) using the dry-weight assay. The dryweight cell assay is not sensitive enough to measure the initial cell concentration. For the purposes of the electron balance, the initial cell concentration may be assumed to be zero.
Theoretical Analysis Batch fermentations are are autocatalytic in nature. As the reaction proceeds, the concentration of the catalyst (i.e., cells), cells), increases exponentially. The rate equation for cell growth is
dx dt
= µ x
(2)
where x is the cell concentration and t is time. The specific growth rate (µ), which varies with temperature, pH, and the substrate and product concentrations, is constant early in the fermentation. As a result, the growth is exponential. However, as substrate concentration decreases, or inhibitory products accumulate to a significant level, µ begins to drop. As the cells enter the stationary stationary phase, µ equals zero.
When µ is constant, Equation (2) can be integrated to yield
ln(x)= µ t +c
(3)
where c is an integration integration constant. Equation (3) indicates that a plot of ln(x) versus t should be linear with a slope slope equal to the specific growth growth rate. Correlation coefficients for a linear fit of Equation (3) to Z. to Z. mobilis growth data higher than 0.99 are not uncommon over a several-hour period (Worden, 1982). The mass of product per unit mass of substrate is given by the yield coefficient, coe fficient, Y p/s. The slope of a plot of the p roduct concentration versus the substrate concentration gives the yield coefficient Y p/s. Alternatively, Y p/s may be determined using substrate and product concentrations from any two times times during the fermentation. When using this latter method, the changes in the substrate and product concentrations should be as large as possible to minimize the effect of errors in the assays. Thus, initial and final concentrations are good choices. An electron balance can be used to evaluate the relative chemical-energy contents of the fermentation products and substrates and to determine the reaction efficiency. The electron balance entails comparing the total number of available electron equivalents contained in the substrate consumed to those contained in the products formed. formed. The balance can be performed using initial and final concentration data. The electron balance can be shown to be a linear combination of elemental mass balances. Electron balances are calculated from reductance degrees of the substrates and products. The reductance degree (γ), of a compound is defined as the number of equivalents of electrons available for transfer to molecular oxygen during combustion, on a C-mole basis. One C-mole is the mass mass of the substance containing 1 mole (12 g) of carbon. Reductance degrees are calculated using using the electron valences as the of the number of available electrons for each element: 4, 1, -2, and -3 for carbon, hydrogen, oxygen, and nitrogen, respectively. The reductance degree of the glucose (C6H12O6) is calculated below.
6 C 4 eγ = mole C
6 O - 2 e- mole 4 e+ + ] = mole H mole O 6 Cmol Cmol 12 H 1 e-
-
where e = electron equivalent C = carbon equivalent H = hydrogen equivalent O = oxygen equivalent
The elemental composition of cell mass is almost almost constant among species. A representative formula that can be used for Z. Z. mobilis is CH1.8O0.5 N0.2. This formula indicates that 1 C-mole of Z. Z. mobilis cells is 24.6 grams. Table 1 shows a sample Table 1. Available Electrons in Fermentation Products
Ethanol Cells
Product Yield 0.48 0.032
Yield (C-mole) 0.209 0.0013
Total Electron Equivalents
ϒ 6 4.2
Electron Equivalents 0.125 0.005 0.13
electron-balance calculation for a set of experimental data. The cell and ethanol yields are expressed in g product produced per g glucose consumed. Ammonia and carbon dioxide are not considered in this balance, because their reductance degrees are zero. Yeast extract is ignored in the balance, because Z. because Z. mobilis does not metabolize carbon sources within the yeast extract to a significant degree. Since 1 gram of glucose contains 0.133 electron equivalents, the electron balance for this example would be within 2% of closure. Students should verify the calculations above. The electron balance can also be interpreted as a type of energy balance, because heats of combustion are directly proportional to reductance degrees for a wide range of organic materials, such as alcohols, sugars, alkanes and biomass. The energy-recovery efficiency of the fermentation can be evaluated using using the enthalpy of the reactants and products. In the example above, 94% of the available electrons were recovered in the ethanol, and only 4% in the biomass.
Symbols
γ = reductance degree µ = specific growth rate c = integration constant e = electron equivalent t = time x = cell concentration YEtOH/glucose = ethanol yield coefficient Y p/s = general product yield coefficient
References Bailey, J.E. and Ollis, D.F., Biochemical D.F., Biochemical Engineering Fundamentals, Fundamentals, pp. 100-130, McGraw-Hill Book Co., New York, pp. 757-758, 1986. Boyer, Rodney F.(1986) Modern F.(1986) Modern Experimental Biochemistry, Biochemistry, pp. 277-278, Addison Wesley Pub. Co., New York. Lee, K. J., M. L. Skotnicki, D. E. Tribe, and P.L. Rodgers, “The Effect of Temperature on the Kinetics of Ethanol Production by Strains of Zymomonas Zymomonas mobilis”, mobilis”, Biotechnology Biotechnology Letters, Letters, vol 3, No. 6, pp. 291-296 (1981). Worden, R. M., “A Kinetic Study of Ethanol Production by Zymomonas by Zymomonas mobilis,” mobilis,” M.S. thesis, University of Tennessee, Knoxville (1982).
Appendix 1: Details of of Experimental Procedure I.
First Lab Period A. Preparation of Growth Media 1.
2.
3.
4. 5. 6.
B.
Prepare 500 mL of 2% yeast extract and 4% peptone in in a 1 L flask. (The solution should be twice as concentrated as desired in the fermenter.) Stopper the flask with a foam plug. Prepare 500 mL of 4% glucose solution in a second 1 L flask apparatus. (The glucose should be twice as concentrated as desired in the fermenter.) Stopper the flask with a foam plug. Prepare two 250 mL “inoculum-culture” flasks, each containing 100 mL of medium with the following following composition: 5% glucose, 1% yeast extract. Cap with a foam plug. Remove the top of the fermenter, and check that the interior components are clean. Replace the top, carefully seating the top against the o-ring seal. Hand tighten the top. After inquiring whether other students may need to autoclave anything, autoclave the reactor, all flasks containing medium, and several screw-top sampling vials. After autoclaving, store store the various solutions at room room temperature. Do not add the medium components to the bioreactor until it is time to inoculate the bioreactor. The solutions will remain remain sterile indefinitely, but opening one of them may lead to contamination by airborne microbes. Minor contamination at the time the bioreactor is inoculated is inconsequential, because there would be many orders of magnitude more S. cervisiae cells added than contaminants, and the fermentation would be over long before the contaminant microbes could grow to a substantial concentration.
Preparation of the Fermenter Vessel for Autoclaving
1. 2.
3. 4. 5. 6. 7.
II.
Make sure sure the power to the unit is off and the water supply to the back of the fermenter is turned off. Remove the motor drive from the top of the vessel, and place it on the motor mount on top of the cabinet. Place the stainless steel autoclaving cap over the bearings where the motor rested. Disconnect all probes and cables from the fermenter. Disconnect the air line so that the sterile filter remains with the fermenter vessel. Disconnect the black water lines from the base of the fermenter. Remove the sampler bulb and insert cotton into the bulb port. Close the valve on the sampler assembly. Any openings through which microbes could enter should be covered or filled with cotton and then wrapped wrapped in aluminum foil. foil. Masking tape can be used to hold the aluminum foil in place. Make sure the feed tube of the fermenter is clamped and stoppered with cotton and wrapped in aluminum foil.
Note: If liquid medium is being autoclaved in the bioreactor vessel, any tubes that are submerged in the liquid medium should be sealed to prevent loss of medium during pressurization; silicone tubing attached to the top of the tube can be clamped to prevent loss of liquid during autoclaving. Use a small piece of silicone tubing to connect the acid and base ports as shown in Figure 2. (Silicone tubing can withstand the heat of autoclaving, while Tygon tubing will melt). Steam must be able to freely flow through a sterile filter or cotton plug into and out of the headspace of the bioreactor. Otherwise, the vessel will shatter during pressurization or depressurization. If you are unsure whether ventilation is adequate, ask the instructor. 8. Compare the autoclaving setup to that in Figure 2. If in doubt, check with the instructor prior to to autoclaving. Several glass sample bottles bottles that fit the sampling assembly should be autoclaved, with their lids loose enough to allow pressure equilibration. equilibration. A large glass funnel should be wrapped wrapped in paper and autoclaved. The funnel will be used to transfer the medium components into the fermenter. 9. Label materials to be autoclaved with the group letter and date to be used to avoid misunderstandings with students in the other section. 10. After asking whether anything else needs to be autoclaved, load and start the autoclave. Detailed instructions for for using the autoclave are provided in another section of this manual. If you have questions regarding use of the autoclave, please ask the instructor. 11. Schedule an appointment with the teaching assistant assistant to do the preparation described below. It is necessary do this preparation at the prescribed times to obtain good data during the second lab period . Preparation for Day Two
A. Preparation of the Inoculum Culture (4 (4 PM the day before the second lab period) 1.
2.
3.
B.
Inoculation of the Fermenter (early morning (8 (8 AM) before Second Lab Period) 1. 2.
3.
C.
Seed the two inoculum flasks with refrigerated brewers’yeast “working culture” about 12 to 15 hours before the bioreactor is to be inoculated. inoculated. For example, if the bioreactor is to be inoculated at 8 a.m. Tuesday, the inoculum culture should be seeded late Monday afternoon. Seed one of the inoculum flasks with about 5 mL of working culture, and the other with about 0.5 mL. The second culture will reach maturity maturity several hours later than the first. Thus, one of the flasks should have a culture at the proper stage of development (mid-exponential phase) when it is time to inoculate the reactor. Details of culture-transfer technique are are given in another section of this manual. Return the working culture to the refrigerator, and incubate the two inoculum culture flasks at 30 to 35°C in the large incubator. Shaking is not needed. The inoculum culture is sufficiently mature when it is highly turbid and foams when the flask is swirled. New cultures usually mature mature within 10 to 20 hours after inoculation, depending on the growth temperature, the amount of inoculum added, and the age of the refrigerated working culture. If both inoculum cultures are mature, use the one that was seeded with 0.5 mL of working culture.
Without turning the power on, install the bioreactor vessel on the console. Place the motor assembly on top of the vessel. Connect the two black water lines to the base of the fermenter. Place the temperature-sensing rod into its slot in the headplate. Turn on the water supply to the heat exchanger at the back of the fermenter. Using the sterile funnel, pour the glucose and yeast extract solutions into the fermenter.
Turn the POWER switch on, and input the setpoints for the temperature and agitation rate. Suggested ranges are are 30 to 35°C and 50 to 200 rpm, respectively. Setpoints are changed by turning the SELECTOR knob to the desired variable (pH, temperature, etc.), and turning the MODE MODE knob to SET POINT. The setpoint value is set with with the INC/DEC button. Once the proper value is shown, setting the MODE knob to CONTROL will engage the controller and show the measured value of that variable. 1.
Withdraw approximately 5 mL of sterile medium into Eppendorf centrifuge tubes using a 5 ml syringe attached to the sampler assembly. Expel any liquid trapped in the sampler tube by squeezing the suction bulb before drawing a sample; this step ensures that the sample has the same composition as the medium in the bioreactor. The sterile medium may be needed later to dilute fermentation-broth samples for the turbidity assay. Also, the absorbance of the sterile medium must be measured to correct
the turbidity data for absorbance of the medium. Refrigerate the medium until needed, to avoid possible microbial growth. 2.
When the bioreactor has reached its setpoint temperature, inoculate the bioreactor by adding an appropriate amount of inoculum culture (usually about 5% of the total liquid volume in the bioreactor).
D. Withdraw an initial initial sample immediately after inoculation using the sampling sampling assembly and a clean, dry tube. About five mL should be sufficient sufficient to perform glucose, ethanol and cell turbidity assays. Properly label and refrigerate refrigerate the sample for processing processing during the second second lab period. During the second period, remove the cells from a portion of this sample; then label and refrigerate the clarified sample for glucose and ethanol assays during the third lab period. E.
Turn on the colorimeter (or spectrophotometer) spectrophotometer) so that it is warmed up when you are ready to make the first first turbidity measurement. Specific instructions on the use of these instruments are found in another section of this manual.
III Second Lab Period A. Process the initial sample taken just just after inoculation. 1. 2.
B.
Warm the sample to room temperature and measure its OD (600 nm) vs. a distilled water blank. Remove the cells from a portion of the initial sample and refrigerate the clarified medium in a capped Eppendorf tube for glucose and ethanol assays during the third period. To do this, place equal volumes of fermentation broth into into each of two 1.5 mL Eppendorf tubes. tubes. Cap the tubes, and place them into slots diametrically opposed from one another in the rotor; this arrangement keeps the rotor balanced. Put the lid on the rotor, and tighten the knob to lock the lid in place. The rotor must be well balanced, with the lid on securely during operation to prevent damage to the centrifuge. If the centrifuge vibrates excessively on start-up, immediately turn it off, and consult the instructor. Close the top, and hold down the black button to activate the centrifuge. When the centrifuge reaches full speed, release the button. After the rotor has come to a complete stop, stop, the top may may be opened. The cells should have settled into a “pellet” at the bottom of the tube, leaving a clear supernatant. Transfer about 1 mL of the supernatant to another, labeled Eppendorf tube, and store it in the refrigerator.
Measure the OD (600 nm) of a 3 mL sample about every 20 min, and measure its OD (600 nm) vs. a distilled water blank. The OD of the cell suspension suspension is directly proportional to the cell concentration up to an OD value of about 1. Therefore, if the OD is greater than 1.0, dilute quantitatively with warm, sterile growth medium to to bring the OD value below 1. When analyzing the data, simply multiply the OD value by the dilution factor to put all values on an equal
basis. Keep a running plot of ln(OD) vs. elapsed time to determine whether the culture is exhibiting exponential growth. C.
(Optional) At regular intervals (about 45 to 60 min), take a small sample, remove the cells by microcentrifuging, and save about 1 mL of the clarified liquid medium in a capped, labelled Eppendorf tube.
D.
Learn how to perform the glucose and ethanol assays by studying the instruction sheets that come with the assay kits. Experience has shown that such early preparation is important. Errors made in developing the ca libration curves or proper dilution of the samples prior to performing the assays have resulted in students getting no useful ethanol and glucose data. If there are questions, please contact the instructor or teaching assistant before the third lab period. (Key points to keep in mind are (1) some samples may be too concentrated to keep within the linear range of the assay and will therefore need to be diluted into the linear concentration range; estimate what dilution (if any) will be needed for each of the samples. (2) To convert the OD resulting from the chemical reaction of the assay into concentration units, a calibration curve is required. The calibration curve is made by plotting the OD of at least two samples having a known concentration (the standards) as a function of c oncentration. Students often have to make up these standards. Good choices for standard concentrations are zero and the upper value of the linear range. That way, the calibration curve can be simply a straight line drawn between the two points. All samples should be diluted so that their concentrations lie between the concentrations of the two standards.
IV Day Following Following Second Lab Period (11 AM) A. Taking Final Samples 1.
Take two 25.0 mL samples of the fermentation broth using a volumetric pipette. Place each sample in a 30 mL centrifuge tube, cover the tube, and store it labelled in the refrigerator for dry-weight assays during the third lab period.
2.
B.
Take a final, 1 mL, sample of the fermentation broth. Remove the cells by microcentrifuging and then transferring the liquid medium to a new Eppendorf tube. The tube should be capped, labelled, and refrigerated for glucose and ethanol assays during the third lab period. Shut off the fermenter, and remove the fermentation fermentation vessel from the the console for cleaning during the third lab period.
V Third Lab Period A. Performance of Glucose Glucose Assays Using the Sigma Diagnostic Diagnostic Reagent Kit (Glucose Trinder, Procedure No. 315)
B.
1.
Estimate the dilutions needed to bring all samples into the linear range of the assay: from 0 to 750 mg/dL (7.5 g/L or 0.75%). If the concentrations of the samples are thought to be out of this range, quantitatively dilute them. As an approximate rule of thumb, 1% glucose is consumed for every 0.6 OD increase in the cell concentration during the exponential growth phase.
2.
Prepare the glucose trinder reagent according to the instructions. There may be enough of this reagent left over from a previous group’s experiment.
3.
Set up a series of test tubes labeled for the reagent blank, standard(s), and experimental samples.
4.
Set the spectrophotometer wavelength to 505 nm, and zero it using distilled water in both cuvettes. The procedure for operating the spectrophotometer is given in another section of this manual.
5.
Warm the glucose trinder reagent to room temperature. Pipet 3.0 mL into each tube.
6.
At timed intervals, add 10 µL (0.01 mL) of RO water, standard, or experimental sample to the appropriate tube and mix. The reagent blank receives 10 µL of RO water.
7.
Incubate each tube for exactly 18 minutes at room temperature. Then, transfer the contents to the sample cuvette, and measure the absorbance at 505 nm.
8.
Subtract the absorbance of the reagent blank from the absorbance of each experimental sample and standard to obtain the absorbance due to glucose.
Performance of Ethanol Assays with Sigma Diagnostic Diagnostic Kit (Ethanol, Procedure Procedure No. 332-UV for Serum) 1.
Follow the method specified for conducting the assay in serum. Estimate the dilutions needed to bring all samples into the linear range of the assay: from 0 to 300 mg/dL (3.0 g/L or 0.30%). (Alternatively, the OD of the the sample should be below about 1.4 after the color reaction is complete). complete). If the concentrations of the samples are thought thou ght to be out of this range, dilute them quantitatively. As an approximate rule of thumb, 0.5% ethanol is produced (and 1% glucose is consumed) for every 0.6 OD increase in the cell concentration during the exponential growth phase. The fermentation converts essentially all of the glucose to ethanol upon completion.
2.
Label one ethanol assay vial (stored in the refrigerator) for each sample to be assayed. Typically, one reagent blank, two standards, and two to five experimental samples will be assayed. Add 3.0 mL of Glycine Buffer
Reagent (stored in the refrigerator) refrigerator) to each vial. Each assay costs several dollars, so please do not use more than needed. An ethanol standard is included with the reagent kit. Note: test tube or other containers cannot be substituted for the assay vials; even though the vials may appear empty, they contain a small amount of dried reagents(enzyme and NADH) that are necessary for the assay to work. 3.
Cap and invert each vial gently several times. Do not shake, as shaking denatures the enzyme.
4.
Set the spectrophotometer wavelength to 340 nm, and zero using distilled water in both cuvettes. As 340 nm is on the fringe of the UV spectrum, spectrum, cuvettes suitable for UV detection should be used.
5.
At timed intervals, add 10 µL (0.01 mL) of RO water, standard, or experimental sample to the appropriate vial, and mix by gentle inversion.
6.
Incubate each tube for 10 minutes at room temperature. Then, transfer the contents to the sample cuvette, and measure the absorbance at 340 nm.
7.
Subtract the absorbance of the reagent blank from the absorbance of each standard and experimental sample to obtain the absorbance due to ethanol.
8.
If one standard is used, the ethanol concentration of the unknowns can be calculated using the formula shown below: A sample - Ablank x Concentrat ion of Standard A standard - Ablank
9.
(5)
Alternatively, a series of known ethanol standards may be run, and a calibration curve may be developed.
C. Dry-Weight assay 1. 2.
3.
Withdraw two 25.0 mL samples of fermentation broth using a volumetric pipette. Place each sample in a 30 mL centrifuge tube kept on ice. Both of the tubes must weigh the same to balance the rotor of the centrifuge. Weigh them, and add enough RO water to to the lighter tube to equalize the weights. Place the two tubes in diametrically opposite slots in the rotor of the Sorvall refrigerated centrifuge; this arrangement keeps the rotor balanced. Put the lid on the rotor, and tighten both knobs (the larger one first) to lock the lid in place. The rotor must be well balanced, with the lid on securely during operation to prevent damage to the centrifuge. If the centrifuge vibrates excessively on start-up, immediately turn it off, and consult the instructor.
4.
Close the top, and centrifuge at 8000 rpm for 10 min. When the rotor has fully stopped, the top may be opened. The cells will have settled settled into a “pellet” at the bottom of the tube.
5.
Label and refrigerate a sample of the supernatant (about 1 mL) for glucose and ethanol assays. Remove and discard the remaining supernatant with a pipette, taking care not to lose any of the cell mass.
6.
Resuspend the cell pellet in about 20 mL of RO water and centrifuge again, as before. This water wash removes residual residual traces of the medium medium from the cells.
7.
Repeat the RO water wash.
8.
Weigh an aluminum weighing pan on the analytical balance. Initials may be engraved into the aluminum with a sharp object.
9.
Transfer all of the cell mass into the weighing pan using a small amount of RO water. A Pasteur pipette is useful for resuspending the cells and transferring the suspension to the pan.
10. Dry the sample overnight at 95°C in the drying oven. The oven should be set to about 3, with the vent slightly open. VI. Day after Third Lab Period (11 AM) 1.
Once the sample is dry, cool the pan in a desiccator to prevent water absorption from the air.
2.
Weigh the pan on the analytical balance, and calculate the mass of dry cells by difference.
Figure 1
Setup for Autoclaving and Aseptic Transfer of Medium for Autoclaving
Enzyme Kinetics Introduction Enzymes are biological catalysts composed of protein. Catalysts increase the rate of a chemical reaction by lowering the activation energy. They do not shift the equilibrium equilibrium of the reaction, nor are they consumed in the reaction. Enzymes often exhibit greater specificity than other types of catalysts. Many can distinguish between very similar similar substrates (e.g., optical isomers) isomers) and produce optically pure (chiral) (chiral) products. For this reason, enzymes are useful in the production of pharmaceuticals and other fine chemicals, ch emicals, particularly those with undesirable stereoisomers. The catalytic efficiency of an enzyme is determined by measuring its reaction kinetics. The objectives of this experiment experiment are to measure the reaction kinetics of the enzyme e nzyme fumarase and to determine the mechanism by which enzyme inhibitors affect the enzyme.
Background Michaelis and Menten recognized that the effect of substrate concentration on the reaction rate varied with the the relative concentration of the substrate. substrate. At low concentrations, enzymes exhibited first-order behavior with respect to substrate concentration, but at high high concentrations, they exhibited exhibited zero-order behavior. Michaelis and Menten developed a two-step mechanistic model predicting that the enzyme and substrate first combine reversibly into a complex, and then the complex react irreversibly to form the product and regenerate regenerate the free enzyme. If the concentration of the complex is assumed to be either at pseudo-steady-state or in equilibrium, the rate equations for the two steps lead to Michaelis-Menten model:
v=
where
dS dP V max S == dt dt K m + S
(1)
v = reaction velocity (rate) Vmax = maximum reaction velocity K m = half-saturation or Michaelis constant S = substrate concentration P = product concentration t = time
The hyperbolic form of this model is able a ble to predict both the zero-order and a nd first-order regimes. Physical explanations for the two regimes regimes are evident from the elementary-step mechanism used to derive the equation. In the first-order regime (i.e., at low S) most of the enzyme molecules are not bound to substrate molecules, and are thus not contributing to the reaction rate. As the substrate concentration is increased, the concentration of enzyme-substrate complexes increases, and hence the reaction rate, increases
proportionally. In the zero-order regime (i.e., at high S), essentially all of the enzyme molecules are already bound to substrate molecules. Thus, further increases in the substrate concentration do not increase the reaction rate. This explanation allows physical significance to be given to each of the two kinetic constants: Vmax represents the reaction rate that would theoretically occur at an infinitely high substrate concentration, and K m is the substrate concentration that produces half the maximum reaction rate. The “affinity” of the enzyme for the substrate is inversely proportional to the K m. Hence, the lower the affinity, the higher higher the S value needed to give half the maximum reaction rate. Both K m and Vmax can be determined from experimental measurements of v as a function of S. The enzyme fumarase adds water to the double bond of fumarate, converting it to Lmalate, as shown in Figure 1. Fumarase is a stereospecific enzyme; only the trans configuration of fumarate will react with the enzyme, and L-malate is the sole product (Dixon, 1979). Both fumarate and malate are intermediates intermediates in the Citric Acid Cycle.
Figure 1. Reaction Catalyzed by the Enzyme Fumarase Acids such as glycine, malate, citrate, adipate, trans-aconitate, glutarate, succinate, and malonate can act as inhibitors of fumarase. Phosphate and citrate are activators of fumarase, (i.e., they increase the reaction rate), while sodium chloride is an inhibitor. The reaction rate may be monitored by assaying (measuring) the concentration of either the substrate or product as a function of time. Because double bonds absorb light at 300 nm, the fumarate concentration can be conveniently assayed by its optical density (OD), also referred to as absorbance, at this wavelength.
Experimental Equipment and Procedure The kinetics of the conversion of fumarate to L-malate by fumarase are measured by monitoring the OD at 300 nm using a Perkin-Elmer Lambda 3B spectrophotometer. The data are recorded and analyzed using a PC-based, automatic data acquisition system capable of recording the OD of standards, internally calculating a calibration curve, and then converting subsequent OD data directly into concentration units. It can also numerically differentiate the data over a user-specified time period to give the initial reaction rate.
Details of the use of the Lambda 3B spectrophotometer and data acquisition system are given in the Appendix. The Lambda 3B is configured to be controlled by the PerkinElmer PECSS software located on the PC. Thus, setting the wavelength, zeroing the instrument, starting and stopping the data acquisition, etc. should all be done using the computer. During the first laboratory period, students should prepare the solutions needed for the kinetic assays and develop a calibration curve giving the OD (300 nm) vs. fumarate concentration. The key capabilities of the PECSS software, software, such as doing time-drive time-drive runs, determining the reaction rate using slope calculations, inputting and deleting data for a calibration curve, and storing and recalling data from the hard disk should be learned during the first period. In this way, the second and third third periods may be most effectively used measuring reaction rates. rates. A table giving suggested dilutions to make the fumarate solutions is provided in the Appendix. Adjustable, automatic pipettors are available to accurately and quickly quick ly transfer small volumes of liquid for preparing solutions. To adjust the volume dispensed, dispensed, gently pull the plunger out and turn. Chemical reaction rates are temperature dependent. To obtain consistent results, all solutions (except the enzyme) should be at room temperature when the assays are performed. It may be useful to arrange with the teaching assistant for the appropriate solutions to be taken out of the refrigerator an hour early to warm up. Enzymes spontaneously denature (lose activity) at a rate that increases with temperature. Thus, the enzyme concentrate should be kept either refrigerated (not frozen) or on ice at all times. The small volume of enzyme concentrate will will warm up to the reaction temperature instantaneously when added to the cuvette to initiate the reaction. It is good practice to calculate the volume of enzyme concentrate needed for the day’s experiments and transfer just that amount to a test tube that is kept on ice. If any of this concentrate is left when the experiments are complete, do not return it to to the original bottle. This precaution minimizes the risk of contaminating the stock concentrate solution. Also, because there are two lab sections that use the same reagents, plan the experiments so that data would not be wasted if something were to happen to the bottle of enzyme concentrate between the second and third periods. For instance, make each set of runs runs conducted during a lab period internally complete. Alternatively, at the end of the the second lab period, calculate how much enzyme concentrate will be needed for the third lab period, and store this amount in a capped, labeled plastic Eppendorf test tube in the refrigerator. Reaction rates are also a function of pH; if the pH is outside of a certain range, the enzyme will denature. The substrate and inhibitor solutions solutions should be titrated to a pH of 7.0 before use. During the second and third periods, p eriods, the enzyme assays will be conducted condu cted for different concentrations of fumarate, and, in some cases, ca ses, in the presence of citrate or phosphate. pho sphate. Each assay takes only a few minutes, because only the initial, linear portion of the curve is needed. Thus, a large number of experimental runs may be conducted. The more data obtained, the more accurately the kinetic constants will be determined.
Theoretical Analysis The Beer-Lambert law, shown below, indicates that relates the OD of a dissolved compound is directly proportional to its concentration:
i OD = log( ) = a b c io where
(2)
OD = optical density io = intensity of the incident light i = intensity of the transmitted light a = extinction coefficient b = length of the light path c = concentration of the absorbing substance
In spectrophotometric assays, it is necessary to correct for light absorbance by substances other than the compound of interest. A “blank” is used for this purpose. The blank is identical to the unknown samples, except it has none of the compound being assayed. Thus, subtracting the OD of the blank from the OD of the samples gives the OD due exclusively to the compound of interest. Initial velocities (reaction rates) of enzymatic reactions are determined using the following dynamic version of the Beer-Lambert:
v=
where
( ∆c)
∆t
=
( ∆OD) ( ∆t)(a)(b)
=
( ∆OD)(chart speed) (distance)(a)(b)
v = reaction velocity t = time elapsed a = extinction coefficient b = length of the light path (cuvette width) c = concentration of the absorbing substance OD = optical density
The extinction coefficient, which varies with the wavelength, is either determined experimentally using known standards or obtained from the literature. literature. The path length is 1 cm for most cuvettes. The experimental method entails quickly mixing mixing the enzyme with the substrate solution in the cuvette, placing the cuvette into the spectrophotometer, and recording the OD as a function function of time. The reaction rate is then determined determined from the slope of the OD-vs.-time OD-vs.-time curve. The initial rate is characteristic of the initial initial substrate concentration in the mixture. Once the initial reaction rate has been measured for a variety of substrate concentrations, an appropriate kinetic model is fit to the data. Graphical and numerical methods have been developed to evaluate K m and Vmax from experimental reaction-rate data. The Michaelis-Menten equation may be rearranged into into
(3)
linear forms, such as the Lineweaver-Burk and the Hanes-Woolf forms, that facilitate the graphical approach (Segel, 1986; Robyt and White, 1986). The Lineweaver-Burk equation, shown below, is obtained by inverting both sides of the Michaelis-Menten equation and rearranging:
1 v
=
1 K m 1 + V max S V max
(4)
This equation indicates that if 1/S were plotted vs. 1/v, a straight line should result with an intercept of 1/Vmax, and a slope of K m/Vmax. The Hanes-Woolf equation is obtained by dividing both sides of the Michaelis-Menten equation by S before inverting and rearranging:
S v
=
1 V max
S +
K m V max
(5)
In this case, a plot of S/v vs. S should give and intercept of K m/Vmax and a slope of 1/Vmax. The values of the kinetic constants determined using the Hanes-Woolf plot may differ slightly from those using the Lineweaver-Burk plot. The Michaelis-Menten model cannot predict the the effects of enzyme inhibitors. inhibitors. More complex models have been derived for this purpose based on elementary-step mechanisms that include usually interactions between the enzyme and inhibitor molecules. These models typically have the same same general form as the Michaelis-Menten Michaelis-Menten equation, except that Vmax or K m is a function of the inhibitor concentration, instead of being a true constant. Two classic examples are the competitive and noncompetitive inhibition models. In competitive inhibition, the substrate substrate and inhibitor molecules both bind to the enzyme molecule at the active site. Thus, the inhibitor “competes” with the substrate for that site. The kinetic model derived for this case is shown below:
v=
V max S S + K s (1 + I/ K I )
In this equation, the inhibitor apparently reduces the affinity of the enzyme for the substrate. The higher the value of [I], [I], the higher the apparent K m, and the lower the apparent affinity. In noncompetitive inhibition, the inhibitor inhibitor is assumed to bind at a site other than the active site. The resulting kinetic model is given below:
(6)
V max S 1 + I/ K I v= K m + S In this case, the inhibitor acts by decreasing dec reasing the apparent maximum reaction rate. Because competitive and noncompetitive inhibitors affect the observed enzyme kinetics in different ways, these two modes of inhibition can be distinguished experimentally by determining the effect of the inhibitor on the apparent K m and Vmax. The Lineweaver-Burk and Hanes-Woolf plots are well suited for for this purpose. Graphs summarizing the effects of various types of inhibitors on these plots are given by Bailey and Ollis (1986).
Symbols a = extinction coefficient b = length of the light path (cuvette width) c = concentration in moles i = intensity of the transmitted light io = intensity of the incident light I = inhibitor concentration K I = inhibition constant K m = half saturation or Michaelis constant K m,app m,app = apparent Michaelis constant OD = optical density P = product concentration S = substrate concentration t = time v = reaction velocity Vmax = maximum reaction velocity Vmax,app = apparent maximum reaction velocity
References Bailey, J.E. and Ollis, D.F.(1986) Biochemical D.F.(1986) Biochemical Engineering Fundamentals, Fundamentals, pp. 100-130, McGraw-Hill Book Co., New York. Bock, Robert M., and Robert A. Alberty, “Studies of the Enzyme Fumarase. I. Kinetics and Equilibrium”, J. Equilibrium”, J. Amer. Chem. Soc., Soc., 75, p. 1971, 1953. Boyer, Rodney F.(1986) Modern F.(1986) Modern Experimental Biochemistry, Experimental Biochemistry, pp. 277-278, Addison Wesley Pub. Co., New York. rd
Dixon, Malcolm, Enzymes Malcolm, Enzymes,, 3 ed., Academic Press, New York, NY, 1979, pp. 229, 264265, 316-318,396.
(7)
Massey, V., “Studies on Fumarase 2. The Th e Effects of Inorganic Anions of Fumarase Activity”, Biochem. Activity”, Biochem. J., J., 53, p.67, 1952. Massey, V., “Studies on Fumarase 3. The Effects of Temperature”, Biochem. Temperature”, Biochem. J., J., 53, p.72, 1953. Massey, V., “Studies on Fumarase 4. The Th e Effects of Inhibitors on Fumarase Activity”, Biochem. J., J., 55, p.72, 1953. Robyt, J.F. and White, B.J., Biochemistry B.J., Biochemistry Techniques Theory and Practice, Practice, Brooks/Cole Pub. Co., Prospect Heights, Il., pp. 44-5, 1986. Segel, I.H., Biochemical I.H., Biochemical Calculations, Calculations, 2 291, 1976.
Appendix 1:
nd
ed., J. Wiley & Sons, Inc., New York, pp. 290-
Equipment and reagents needed
fumarate fumarase L-malate citric acid spectrophotometer and recorder UV-grade polymethacrylate cuvettes pH calibration solutions pH meter potassium phosphate (monobasic) volumetric flasks gloves Parafilm pipettes (10µL, 20 µL, 2 mL, and 5 mL) 0.1 M NaOH
Appendix 2: Details of of Experimental Procedure I
Preparation of Stock Fumarate, Citrate, and Phosphate Solutions A. Prepare 500 mL of 18.75 mM mM fumaric acid, 50 mL of 42 mM mM citric acid, and 50 mL of 400 mM potassium phosphate (monobasic). B.
Adjust the pH of the the three solutions to 7.0 using NaOH solution. solution. It is good practice to dissolve the acid in about 90% of the total volume of water, titrate to the proper pH, and then add more water to obtain the desired total volume. The pK values of fumaric acid are 3.0 and 4.4. Thus, fumarate buffers strongly between up to about 5.5, and weakly near neutral pH. To titrate to pH 7.0 quickly and without overshooting, it may be convenient to use a more concentrated NaOH solution below pH 5.5 and a more dilute one above 5.5. Citric acid and phosphate have pK values of 6.4 and 7.2, respectively, and thus be less likely to overshoot.
C.
Prepare diluted stock solutions from the original stock solution. Table 5 gives sample calculations of convenient dilutions. The left hand column gives the final concentration of fumarate in the cuvette after 2 mL of the diluted stock
solution has been combined with 1 mL of distilled water and 10 µL of fumarase enzyme. The second column gives the diluted stock solution concentration made by combining the volumes of original stock solution and distilled water shown in columns 5 and 6. Experience has shown that the most dilute samples often produce poor results, so this concentration may be omitted. Check the pH of the diluted stock solutions, and adjust if necessary. D. Because fumarate fumarate is a common metabolite, it makes an excellent growth substrate for microbes. Thus, all solutions should be refrigerated when not in use to avoid microbial microbial growth. However, all solutions (except the enzyme) should be warmed to room temperature before making dilutions or conducting assays, because reaction rates rates are strongly strongly affected by temperature. temperature. Since it takes a while to warm the solutions, it is advisable to have someone re move the solutions from the refrigerator about an hour before the lab period is to begin. The microwave can be used to warm the solutions more rapidly.
Table 1. Dilution Schedule for the Fumarate Solutions
II.
Final Curvette Conc for 2:1 ratio (mM)
Diluted Stock Conc (mM)
% stock in solution
volume made (mL)
stock added (mL)
water added (mL)
# of runs availabl e
12.45847
18.75 18.75
100
500
500
0
79
9.966777
15
80
100
80
20
50
7.475083
11.25
60
100
60
40
50
4.983389
7.5
40
100
40
60
50
2.865449
4.3125
23
100
23
77
50
2.491694
3.75
20
100
20
80
50
1.245847
1.875
10
100
10
90
50
0.622924
0.9375
5
100
5
95
50
0.498339
0.75
4
100
4
96
50
Instrument and Computer Startup (Pages (Pages 3-6 of the the Perkin-Elmer Perkin-Elmer PECSS manual) A. Remove any cuvettes from the sample and reference reference beams. beams.
B.
Turn on the spectrophotometer’s UV bulb using the UV ON switch on the front lower right of the spectrophotometer.
C.
Turn on the main power switch located on the back, right side side of the unit, and allow 30 minutes for the spectrophotometer to warm up.
D. After turning on the spectrophotometer, a calibration must be run. run. To run calibration, press the “safe mem” key on the Perkin Elmer Lambda 3 until the display shows a “C”. Press the “Run” key to start calibration. E.
Allow 10 minutes for calibration. Once machine is finished, an “O” should appear in the display; indicating ready to operate.
F.
Turn on the computer. Anti-virus is out of date, so press “N” at the prompt to say you don’t want to continue with with virus scan. Type “PECSS” at the prompt. The “ready for the next command” prompt should appear.
G. Press SHIFT-F10 to to get the menu. menu. (To get the “Ready for the next command prompt back, press ESC).
III. Autozeroing the Spectrophotometer (Pages 4-11 of the PECSS manual) manual) A. Press SHIFT-F10 to get the menu, and use the arrow keys to move the the cursor to “PARAM”. After ORD type “A” for OD; after speed speed type “120”; after Autosave type “Y”; and after Autoprint type “N”. B.
Insert a cuvette containing the blank solution (RO water for the calibration curve) both the BLANK (rear) position, and the SAMPLE (front) position. Close the lid. Always hold cuvettes by the frosted frosted sides and wipe the clear sides with a Kimwipe before inserting inserting them into the spectrophotometer. Make sure there are no particles or bubbles bubb les on the clear window that would affect the OD readings. Note: only UV-grade cuvettes should be used. Standard cuvettes will absorb too much of the incident light at 300 nm.
C.
Press SHIFT-F10 again, and move the cursor cursor to autozero.
D. Under autozero type “A” after mode (for OD) and “300” after wave (the wavelength in nm). With the cuvettes are in spectrophotometer and the lid closed, type “Y” to begin the autozero procedure. IV. Calibrating the Spectrophotometer (Pages 4-13 to 4-20 of the PECSS manual) A. Press SHIFT-F10 to get the the menu, menu, and choose CALIB. CALIB. Beside method, type a name having up to 8 digits (e.g., (e.g., “groupc”). The calibration-curve data will later be stored under this name. Beside nwave, type “1” to indicate that only one wavelength will be used. Beside curve, type “1” to to indicate that the
calibration curve should be linear and pass through the origin (e.g., have the form concentration = m•OD, where m is the slope). B.
After wavelength w1_, type “300” for 300 nm.
C.
When the the screen displays Change or save the instrument parameters (Y/N)? type “N”.
D. Prepare a standard by mixing two mL of one of the stock solutions solutions described in the table with one mL of RO water. Cap the cuvette with a piece of Parafilm, Parafilm, and gently invert it several times times to mix. Practice mixing quickly without generating many bubbles. E.
Place the cuvette in the SAMPLE holder, and close the lid.
F.
When prompted, enter the concentration of the standard. Press “enter”. Enter a sample identification (e.g., STD01) and then press “Y” beside Ready (Y/N)?
G. Replace the first standard with with the second one. If the first first sample identification ended in two digits, the computer will automatically offer the same name with the number incremented by one for the second sample. Enter the concentration of the second standard and type “Y” under Ready (Y/N)? H. Repeat this process for all all (up (up to 15) standards. standards. After the last last one has been recorded, press ENTER in response to the Ready (Y/N)? prompt. I.
At this point, the computer will calculate a calibration curve and display it on the screen. If the cuvette’s path length length is 1 cm, the slope slope is numerically equal to ⋅ pathlength, as the extinction coefficient in dimensions of OD/concentration pathlength, defined by the Beer-Lambert law. Record the slope and the residual error error (resid err:) for your report.
J.
If mistakes were made in entering the the calibration curve, points points may be deleted with the View/del std option.
K. Choose the Write calib option to have your calibration-curve data written in ASCII format to the file “groupc.cal”. This file can then be printed printed out or copied onto a floppy disk for later analysis. V. Monitoring the Enzymatic Reaction (Pages 4-7 to 4-9 of the PECSS manual) A. Press SHIFT-F10 to get the menu, menu, and choose TDRIVE. Beside reg, type “x”; beside wave type “300”; beside NoPts, type “1000”, and beside int type “0.5” to have a data-logging time interval of 0.5 seconds. After pressing pressing ENTER, you will then be prompted for a minimum and maximum OD (y) values. The default values of 0 and 1.0 should be satisfactory. If not, type in different different values. Also, type in a sample sample identification that is specific for your group (e.g., groupc01). If the name has two digits digits at the end, PECSS will automatically increment the digits by one each run. Do not press ENTER or “y” until the
enzyme and substrate have been combined and the cuvette has been placed in the spectrophotometer. B.
Fill a small ice bucket and with ice. Wear disposable latex gloves when handling the enzyme stock solution to help keep out proteases and other contaminants. With a clean pipette tip, transfer transfer the amount of enzyme needed for the day’s experiments from the original bottle to a 1.5 mL, plastic Eppendorf Eppendo rf tube kept on ice. Tightly seal the original bottle, bottle, and return it to the refrigerator. The fumarate standards, inhibitor solutions and about 100 mL of RO water should be kept at room temperature while in use.
C.
Combine 2 mL of fumarate standard solution with with 1 mL of RO water in the sample cuvette.
D. Rinse the pipet tip used used for the enzyme thoroughly with water. Transfer 10 µL of fumarase from the test tube to the cuvette. Quickly cover the cuvette with a parafilm square, and mix by gently inverting the cuvette several times. Try to minimize bubble formation. E.
Immediately place the the cuvette in the SAMPLE slot of the spectrophotometer, close the lid, and press “y” and ENTER to initiate the data acquisition. The OD data will be plotted in real time on the screen. When the plot is no longer linear, press ESC to stop the run. With experience, the run may be stopped before it is clearly nonlinear without sacrificing accuracy. Depending on the fumerate concentrations, each run takes about 20 to 80 sec.
F.
To leave the TDRIVE TDRIVE menu and analyze the data, type “N” at the Ready (Y/N)? prompt.
VI. Viewing the Data (Page 5-27 of the PECSS manual) A. Press F9 to use the VIEW feature. B.
Type “x” beside reg, and type “a” (for (for automatic) beside min. min. The other slots may be left at the default values. Press ENTER.
C.
Select the time interval over which which to to determine the initial reaction rate. The region should be early enough to be characteristic of the initial substrate concentration, but not so early ea rly that swirling of the fluid inside the cuvette may affect the results (e.g., within within the first couple of seconds). Typically, data in the first couple of seconds should not be used.
D. Although the data can be saved on the hard disk for later analysis, this approach is risky, because the data could accidentally be lost. It is advisable to determine determine the slope (reaction rate) immediately after each run, as described below. VII. Determining the Initial Reaction Rate (Pages 5-19 to 5-20 of the PECSS manual)
A. Press SHIFT-F3 to use the SLOPE feature. B.
VIII.
Type “x” beside reg, and enter the starting starting and ending times for the slope calculation. If the default factor of 1 is used, the calculated slope (designated SL) will be in units of OD/sec. The residual error (designated RE) and linearity factor (designated LF) indicate how well the data conform to a linear linear fit. These values should be recorded, along with the reaction rate, and the time interval used for the slope calculation. The slope value may be divided by the extinction coefficient to calculate the reaction rate. Alternatively, the reciprocal reciprocal of the extinction coefficient may be used as the factor, and PECSS will automatically calculate an enzyme “activity”. However, the calculated activity will have time units of minutes, rather than seconds. Inhibitory Effects of Citrate and Phosphate on Fumarase
A. Autozero the spectrophotometer using using a blank consisting of 2.5 mL water and 0.5 mL inhibitor solution. B.
IX.
Repeat reaction-rate assays using 2 mL of fumarate solution, 0.5 mL of RO water, and 0.5 mL of inhibitor solution. Exiting PECSS (Page 3-7 of the PECSS manual)
A. To temporarily temporarily leave PECSS, use DOS commands, and then return, press ESC to get the Ready for the next command command prompt. Then type “dos” to get the DOS prompt. After the desired DOS commands have been executed, type “exit” to return to PECSS. B.
To quit PECSS, press ESC to get the Ready for the next command prompt, and then type “stop”.
Plasmid Stability Introduction Recent discoveries in recombinant-DNA technology have allowed the insertion of new genetic instructions into living cells. This process allows proteins to be mass produced by microbial species that are well suited to growth in bioreactors. Several pharmaceuticals are commercially produced by genetically engineered organisms. For example, human insulin is now made by genetically modified Escherichia modified Escherichia coli bacteria. The segment of DNA coding for a particular protein is called a gene. Genes can be transferred to a new host cell by a process known as transformation. transformation. Transformation is usually achieved by inserting the gene into a segment of DNA called a vector, and then the vector is transferred transferred into the new host cell. cell. Plasmids are commonly used used as cloning vectors. Plasmids are small small loops of DNA that replicate replicate independently of the rest of the cell. When the cell divides, each daughter cell generally generally receives at least one copy of the plasmid, and thus the plasmid is carried on from generation to generation. However, there is a finite probability that all copies of the plasmid will end up in one of the daughter cells. The other cell, and all its offspring, will lack the plasmid. Because the plasmid-lacking cells consume substrate but do not make the desired product, they reduce the product yield. The effect of plasmid loss loss on reactor productivity can be theoretically theoretically predicted if the probability of plasmid loss during cell division division is known. The objective of this experiment is to measure the probability of plasmid loss during cell division for a recombinant strain of E. E. coli in two different types of growth media.
Background Cloning vectors can be customized to have features well suited for a particular transformation process. Restriction enzymes are used to “cut and paste” the appropriate DNA fragments, sometimes sometimes referred to as cassettes, cassettes, to assemble the vector. Ideally, vectors should have unique restriction sites for several different enzymes (i.e., each enzyme should cleave the vector in only one spot), so the exact location of the insert within the vector is known. They should have an origin of replication for the new host cell, so that the new host cell can process the information information encoded in the DNA. They should have a promoter gene that enables gene expression to be turned on or off to maximize the fermentation productivity. They should also contain at least least one selection marker to allow rapid selection of cells that have successfully successfully taken up the vector. Genes for antibiotic resistance are commonly used as selection markers; cells containing the plasmid will grow in the presence of the antibiotic, while plasmid-free plasmid-free cells will die.
The two most common classes of cloning vectors are viruses and plasmids. Viruses, often called bacteriophages or phages, phag es, are naturally occurring, parasitic entities that attach to a host cell and then inject it with viral viral DNA. This DNA either becomes stably stably integrated into the host’s chromosomes, producing a dormant phase called lysogeny, or it directs the protein-manufacturing capabilities of the cell to produce millions of copies of the virus. This latter phase of the infection, the lytic phase, results results in destruction of the host cell. Lysogenic viruses are used as cloning vectors, because they do not destroy destroy the host cell. Plasmids are double-stranded loops of DNA that frequently contain genes for nonessential cell traits, traits, such as antibiotic resistance. resistance. Methods to transform cells with plasmids generally involve a treatment step that permeabilizes the exterior of the cells. The cells are incubated in a solution containing the plasmid DNA, which is taken up by some of the cells. The transformed cells are then identified by a selection marker marker contained on the plasmid. Plasmids containing several antibiotic-resistance genes are convenient for geneticengineering research, because multiple selection markers can be used simultaneously. An example of such a selection strategy strategy is shown schematically in Figure 1. If a DNA insert is cloned into the ampicillin-resistance gene, as shown in Figure 1b, resistance to that antibiotic will be lost. However, resistance to the other antibiotics will be retained. retained. All cells transformed by the plasmid—both with and without the DNA insert—may first be selected by growth on one of the other antibiotics. Then, these cells can be further screened for resistance to ampicillin; those without ampicillin resistance would contain a DNA insert. The E. The E. coli strain used in this study, EC147, was develop ed by Dr. Pat Oriel of the MSU Department of Microbiology and Public Health using this approach. Once in a suitable host, plasmids are autonomous in the sense that they replicate independently of the rest of the the cellular components. The number of plasmid copies per cell, referred to as the copy number, n umber, varies with the plasmid, the host cell, and the environmental conditions. During cell division, division, each daughter cell normally receives at least one copy of each plasmid. However, there is a finite probability that all the copies will segregate into one of the two daughter cells. The other cell, and all of its its descendants, will not have the plasmid. plasmid. This type of plasmid loss loss is referred to as segregational plasmid instability. instability. Clearly, if plasmid plasmid loss occurs at a significant significant rate during a fermentation where the product is plasmid-encoded, the reactor productivity would be greatly diminished. Replication and expression of the plasmid places an additional metabolic burden on the plasmid-containing cells. Thus, these cells have fewer resources available to fuel growth. As a result, plasmid-containing cells commonly grow more slowly than their plasmid-free counterparts. Moreover, cells typically lose their their plasmids with a higher frequency when grown in relatively poor growth media than when grown in rich media.
Experimental Equipment and Procedure In this experiment, the probability of plasmid loss by E. by E. coli strain EC147 is determined. The specific growth rates of the plasmid-containing and plasmid-lacking cells (µP and µ N, respectively) can be determined during batch growth in LB broth at 35°C. Alternatively,
it may be assumed that µP and µ N are approximately equal; in this case, only the plasmidcontaining cells need to be grown. The specific growth rate is determined by measuring the optical density of the cells during batch growth every 20 minutes and then plotting the data on semi-log coordinates. In addition, the cell concentration and fraction fraction of cells containing the plasmid are measured at the beginning and end of the second lab period. These data are then analyzed analyze d to give the probability of plasmid loss during cell division, as described in the Theoretical Analysis section. There are two ways of distinguishing plasmid-containing cells from plasmid-lacking cells. The plasmid plasmid in E. in E. coli strain EC147 codes for both ampicillin (an an tibiotic) resistance and production of an α-amylase enzyme that degrades starch. If the plasmid plasmid is lost, the cell is no longer resistant to ampicillin and longer makes amylase. To determine whether cells are producing amylase, a properly diluted cell suspension can be plated out on agar plates containing starch. After incubation, the plates can be coated with an iodine stain that darkens in the presence of starch. Colonies having the plasmid will will have a clear halo around them, indicating that the starch starch has been broken down by amylase. Colonies arising from cells without the plasmid will not have the halo. However, experience has shown that as the cells are growing into colonies on the plate, some cells lose the plasmid, and the colony consists of a mixture of plasmid-containing and plasmid-lacking cells. When these colonies are stained, stained, it is often difficult difficult to tell whether there is a halo. Thus, ampicillin may make the best selection marker to determine the fraction of plasmid-containing cells at a given time in the fermentation. The method involved replicate plating, where identical samples of properly diluted cell suspension are plated out on starch/agar plates with and without 50 µg/mL ampicillin. Details of how to make make the ampicillin plates are given in Appendix 2. Only cells with the plasmid should grow on these plates, whereas both cells with and without the plasmid should grow on the plates lacking lacking the ampicillin. The fraction of cells containing the the plasmid can be determined by dividing the number of colonies growing on the ampicillin plates by the number growing on the plates without ampicillin. The iodine-staining method can be used to make sure the cells growing on the ampicillin plates are the desired cells and not some contaminant that happens to be resistant to ampicillin. If the culture is pure, all colonies on the ampicillin ampicillin plate should give halos when stained. However, if the colonies are too close together, their halos will overlap, making it difficult to distinguish which colonies have halos. halos. On the other hand, if the the colonies are too far apart, the total number of colonies counted will be too low to be statistically meaningful. A colony density of between about 30 and 100 cells per plate is optimal. A method has been developed to estimate the the dilution required to give about 100 colonies per plate. Details of the method are given in Appendix 2. The growth flask without antibiotic will be sampled and cells plated once at the beginning of the second lab period and once near the end of the period. Students should take their second sample early enough that the dilutions and plating may be completed by the end of the period. The experiment may be repeated during the third period to explore the effects of alternative growth temperatures or carbon sources (glucose, succinate or glycerol) on plasmid stability. stability. Growth on these alternative carbon sources sources is expected to be much slower than on LB broth. The doubling time on LB broth is about 20 min, while
the doubling time on glycerol is on the order of 1 hour. Because it would be desirable to compare results between media for similar number of cell doublings, the final plating for cells growing more slowly slowly should be done several hours after the end of the lab period. It will be necessary to schedule with the teaching assistant or instructor a time for the lab to be opened for this purpose.
Theoretical Analysis Material-balance and kinetics concepts can be used to derive equations describing the rate of plasmid loss due to segregational instability. The material balance on cells during batch growth is
dx dt
= µ x
(1)
where x is the cell concentration, and µ is the specific specific growth rate. The specific growth rate can be determined by measuring the optical density (OD) of the cell suspension a t 600 nm as a function of time. time. The cell concentration is directly proportional to the OD up to an OD value of 1. As shown by integrating Equation 1, the slope of a semi-log semi-log plot of optical density vs. time should be equal to the specific growth rate. Mass-balance equations analogous to Equation 1 may be written for the plasmidcontaining cells (P) and plasmid-lacking cells (N): dP = µ P (1 - h)P dt dN dt
= µ P (h)P + µ N N
(2)
(3)
where h is the probability that one on e of the daughter cells will not receive the plasmid during cell division, and µP and µ N are the specific growth rates of the plasmid-containing and plasmid-lacking cells, respectively. As described in Chapter 13 of your text, Equation 2 may be integrated, and the resulting expression for P may be substituted into Equation 3. If Equation 3 is then then integrated assuming that at t = 0, N = 0 (i.e., (i.e., that all cells initially have the plasmid), and the solution is put in dimensionless terms, the result is
F n =
1 - α - h 1 - α - h ⋅ 2n( α +h-1)
where Fn is the fraction of cells containing plasmids after n generations, a nd α is the ratio µP/µ N.
(4)
The number of generations (n) can be calculated as shown below if the entire growth process takes place during exponential growth: n=
µ P t
(5)
ln(2)
where t is the duration of growth. Alternatively, n can be calculated from the initial and final total cell concentrations (N + P). The following equation can be derived by integrating integrating Equations 2 and 3 from time t0, when the total cell concentration is (N + P)0 and the fraction of cells containing plasmids is F0:
( N + P ) n( 1 – h) n = F0(1 + h/ β )[2 -2 ]+2 ( N + P )0 α
n
α
(6)
Assuming F0 and α are approximately 1.0, and h is approximately zero, an estimate of n can be found using the expression:
( N + P ) ≅ ( N + P )0
2
n
(7)
Using this approach, n can be estimated based only on the initial and final cell counts and the known dilution factors. Thus, the final cell sample sample could be taken on the morning after the second lab period instead of at the end of the second lab period. In this way, a larger n value would be obtained, and consequently the chances of measuring significant plasmid loss would be increased. The fraction of plasmid-containing cells after n generations may be calculated from the experimental measurements of P and N: F n =
P P + N
The resulting F0 value can then be used in Equation 4 to determine h.
Appendix 1: Experimental Supplies Supplies and Equipment Equipment E. coli strain EC147 petri dishes incubator (4) 250 mL Erlenmeyer flasks for medium preparation 500 mL flask for buffer preparation spreading bar test tubes (2) 1 L flasks for agar preparation 250 mL or 500 mL flask for batch fermentation
(8)
iodine reagent solution (3.0% KI and 0.3% I2 in distilled water) components of LB medium components of M9 buffer alternative carbon sources for M9 growth medium glycerol glucose succinate glutamate agar soluble starch 25 mg/mL ampicillin stock solution 0.5 M MgSO4 stock solution 0.2 or 0.45 µm sterile syringe filters Bunsen burner sterile pipettes wire transfer loop
Appendix 2: Details of of Experimental Procedure I.
Preparation of Growth Media, Agar Plates, and Sterile Test Tubes A. Prepare four 250 mL mL Erlenmeyer flasks of medium: two containing 100 mL each of LB broth, and two containing 100 mL mL of M9 medium. (Either M9glycerol medium or M9-succinate medium may may be used). For both growth media, one of the flasks (the inoculum flask) will be used for growing an inoculum culture, and one (the growth flask) will be used for growth of the cells without antibiotic selection. Also, prepare 300 mL of sterile M9 buffer in a 500 mL Erlenmeyer flask for the serial dilutions. The recipes are given below. Adjust the pH of the solutions to 7.0 with NaOH solution. 1.
LB broth (a rich growth medium): Prepare either a 20 g/L solution of premixed LB medium powder, or use the following individual components: a. 10 g/L tryptone b. 5 g/L yeast extract c. 10 g/L NaCl
2.
M9 buffer : a. 6.4 g/L Na2HPO4⋅7H2O b. 1.5 g/L KH2PO4 c. 2.5 g/L NaCl d. 5.0 g/L NH4Cl
3.
M9 medium (a minimal growth medium): add to the M9 buffer recipe above 4 g/L of one of the alternative carbon sources (glycerol, glucose, glutamate, or succinic acid). Also add enough MgSO4 stock solution to make its concentration 1mM in the medium. Titrate to pH 7.
B.
Prepare enough LB broth with 20 g/L agar and 20 g/L starch for 40 LB-starchagar plates with 50 µg/mL ampicillin and 40 without ampicillin. ampicillin. About 700 mL of LB broth containing agar and starch will be needed for each group of 40 plates. (Neither the starch nor the agar will dissolve until the medium is autoclaved.) This number will allow allow triplicate plating of each dilution at two different times for two different media types, plus a few extras. The ampicillin is added after autoclaving and before pouring the plates.
C.
Prepare about 3 mL mL of a 25 mg/mL ampicillin stock solution. Aliquots of ampicillin stock solution may may be stored frozen in the freezer. freezer. If not, fresh stock stock solution should be prepared. The ampicillin is stored stored in the refrigerator.
D. Obtain 60 test tubes with plastic caps. The test tubes will will be used for serial dilutions of the cell culture prior to plating. E.
Autoclave the the test test tubes and all prepared solutions at 121°C for 15 minutes. Instructions for using the autoclave are given in the final chapter of this manual.
F.
After autoclaving, add ampicillin to the agar for the the ampicillin ampicillin plates as described below. 1.
Cool the agar solution for the ampicillin plates to about 50°C.
2.
Draw the proper volume of 25 mg/mL ampicillin stock solution into a sterile, disposable syringe to give a final concentration of 50 µg/mL. (The proper dilution is 2 mL of 25 mg/mL stock solution per L of medium.) Notice that a small amount of stock solution (about 0.1 mL) is required to fill the sterile filter, so draw a little more into the syringe than is to be added to the flask.
3.
Attach a sterile, disposable filter (0.2 µm pore size) to the syringe tip, and force the desired volume through the filter and into the sterile solution. Flame the neck of the Erlenmeyer flask after removing, and before replacing, the foam plug. Swirl to mix mix the ampicillin throughout the agar.
4.
Pour the agar plates, as described in the Aseptic Techniques chapter of this manual.
5.
Allow the plate to solidify, and leave them overnight without parafilm to allow any water that exudes from the gel to evaporate. Stretch a strip of Parafilm around the edge of each plate to help keep airborne microbes out. Store the plates upside down.
G. Schedule an appointment with the teaching assistant to do the preparation for the second lab period. It is necessary to do this preparation at the prescribed times to obtain good data during the second lab period .
II.
Preparation for the Second Lab Period A. Preparation of the inoculum culture (after (after 4PM the afternoon before the second lab period)
B.
1.
Reserve 5 mL samples of the M9 medium and the LB broth before inoculation. These will be used later as blanks.
2.
Add the proper amount of 25 mg/mL ampicillin stock solution to the two inoculum flasks (one LB broth, one M9 medium) to bring their ampicillin concentrations to 50 µg/mL.
3.
Seed the two inoculation flasks (the ones containing ampicillin) with 5% by volume refrigerated E. refrigerated E. coli EC147 working culture about 15 hours before the flask with no selection is to be inoculated. For example, if the bioreactor were to be inoculated at 8 a.m. Tuesday, the inoculum culture should be seeded late Monday afternoon. Each inoculum flask should be inoculated with a working culture that was grown on the same type of medium. The purpose of growing the inoculum culture in the presence of of ampicillin is to ensure that the fermentation is started with only plasmidcontaining cells (the plasmid containing α-amylase gene has an ampicillin-resistance gene also).
Inoculation of the two growth flasks (early morning morning before the second lab period) 5.
Aseptically transfer about 5 mL of sterile growth medium from each growth flask into a labelled, sterile sterile test tube. Refrigerate the test tubes.
6.
Wash the inoculum-culture cells to remove ampicillin prior to inoculating the growth flasks. Even small concentrations of ampicillin (1-5 µg/mL) can affect the growth of plasmid-lacking p lasmid-lacking cells. a.
For the LB broth inoculum culture, fill and cap four 1.5 mL mL sterile, plastic Eppendorf centrifuge tubes; for the M9 inoculum culture, fill and cap eight tubes. Label the tubes according to type of medium.
b.
Load the 12 Eppendorf tubes into the microcentrifuge so that the rotor is balanced. Each tube should should have another tube located diametrically across the rotor to counterbalance it. Turn on the microcentrifuge for about 20 seconds. After the rotor has stopped, use a sterile pipette tip to remove the supernatant from above the settled cell pellets. Add about 1 mL mL of sterile M9 buffer to each tube and cap the tube. Resuspend the cells using the vortexing machine located on the benchtop near the culture-transfer area.
c.
d. e.
7.
Inoculate the growth growth flasks by pipetting the cells from the Eppendorf tubes to the appropriate flasks.
8.
After inoculation, a 5 mL initial sample should be aseptically transferred from each growth flask to a sterile test tube, labelled and refrigerated. During the second period, these samples will be diluted (if necessary) and plated out to verify that the inoculum contained only plasmid-containing cells.
9.
Incubate the growth flasks at 35°C with 100 rpm shaking.
III. Measuring the specific growth rate of plasmid-containing cells in the two growth media(during the second lab period) A. About every 20 minutes, measure the optical density (600 nm) of a three mL sample of fermentation broth using a distilled water blank. Although sterile media may also be used as a blank, any unexpected cell growth in the blank will affect the results. B.
Maintain a running plot the OD vs. vs. time data on semi-log semi-log coordinates. The slope will be the specific growth rate.
IV. Dilution and Plating of Cell Samples Samples to Quantify Cell Concentration (during (during the second lab period) A. Dilution Procedure to Give About 100 Cells per Agar Plate 1.
Measure the optical density (OD) of a 2 mL sample of each sample sample to be plated at 600 nm using uninoculated growth medium as the blank. If the OD is greater than about 1.5, 1.5 , quantitatively dilute the cell sample with sterile medium until the OD is less than 1.5. Then, back calculate what the OD of the original sample sample would have been. For instance, if the original sample were diluted by a factor of 5 (i.e., 1 part cell culture mixed with 4 parts sterile medium), and the diluted sample had an OD of 0.6, then the OD value of the original sample would be 3.0.
2.
Calculate the required dilution factor using the following equation: Dilution Factor = 100,000 OD x
To continue the example, if the original OD were 3.0, the required dilution to give about 100 cells per plate would be about 300,000. Thus, one volume of the original cell suspension would be diluted with 300,000 volumes of sterile diluent. diluent. Then, 0.1 mL of the resulting diluted cell suspension would be spread out on each plate as described in the chapter on Aseptic Techniques. This method is only approximate, however, so it would be prudent to also plate out a sample ten-fold more concentrated and one ten-fold more more dilute than the one calculated. That is, the 3,000-,
300,000- and 3,000,000-fold dilutions should be plated out. Note that this this dilution prior to plating needs to be done for all samples, regardless of whether the OD of the original sample is less than 1.5. IV. Successive Dilution of Cultures Prior to Plating A. Label several test tubes for each sample with the sample ID and the intended dilution. Aseptically pipet 10.0 mL of sterile sterile diluent into into each tube. B.
Swirl the flask flask containing the culture to resuspend resuspend any settled cells.
C.
Quantitatively transfer transfer an aliquot (e.g., 0.1 mL) of the cell sample to the first test tube using a sterile sterile pipet tip. Mix the sample without wetting wetting the cotton.
D. Using a new pipet tip, quantitatively transfer an aliquot of the diluted sample from the first test tube to the second. E.
Repeat until the proper dilution is achieved.
VI. Spreading Agar Plates for Viable Plate Counting A. Light the Bunsen burner in the culture-transfer area. B.
Place 190 proof (95%) ethanol in a beaker beaker to a depth of about one half inch, and submerge the spreading end of the glass rod in the ethanol. Keep the beaker in the culture-transfer area, but not close to the flame.
C.
Remove the cotton from the mouth of the test tube containing the sample to be diluted, and flame the mouth of the the test tube. Using a sterile pipette tip, withdraw 0.10 mL of diluted cell suspension, trying not to touch the walls of the test tube with the pipetter. Flame the test-tube mouth mouth again, and replace the cotton.
D. Lift the lid of an agar plate, and dispense the cell sample onto the agar at the center of the plate. Replace the lid. E.
Lift the glass rod out of the alcohol and pass the end through through the flame flame to ignite the alcohol remaining on the rod. Warning: ethanol is extremely flammable! Avoid dripping flaming alcohol on anything that might burn. Keep the glass rod level or tilting downward, so the alcohol does not run down toward your hand. Have a plan of action in mind in case of an accidental fire. F.
Once the flame on the glass rod has extinguished itself, itself, cool the spreading portion of the glass rod on the agar away from the cell sample. Use the rod to evenly spread the cell sample over the entire surface of the agar.
G. Return the glass rod to the alcohol. H. Repeat the process for the the other samples to be be plated. When finished, flame flame the glass rod again. I.
Wrap Parafilm around the edge of each of the plates. Invert and label them. Incubate the plates at 35°C overnight.
J.
Schedule with the teaching assistant assistant for someone to remove the plates early the next morning. If the plates are incubated too too long, the amylase clears too large a ring around each colony and makes it difficult to distinguish the staining patterns of adjacent colonies.
K. Schedule a time with the teaching assistant assistant to do the staining and plating described below. VI. Staining plates and counting plasmid-containing plasmid-containing and plasmid-lacking plasmid-lacking colonies (day after the plates have been spread) A. For those dilutions that have appropriate numbers of colonies per plate, count the number of colonies on each plate. Plates with less than than about 20 and more than about 500 should not be counted. It is convenient to to count colonies with the plates upside down. Touch a Sharpie pen to the bottom to mark mark each colony as it is counted. B.
After the counting is finished, finished, apply about 1 mL of the iodine stain to the surface of a plate without ampicillin, and tilt the plate to stain the entire surface.
C.
Allow the stain stain to work five five to ten minutes. If some areas appear to have too little stain, occasionally tilt the plate to redistribute the stain.
D. Count the number of amylase-positive and amylase-negative colonies. In addition to not having the clear halo, colonies without the plasmid also may exhibit a thin, dark ring around their perimeters. The halos may be most easily recognized by holding the plate a few inches above a white sheet of paper while viewing from above. If colonies are too close together to distinguish their staining patterns, simply do not count those cells. Mark each colony you count with a dot on the bottom of the plate below the colony. Use different colors for for the two cell types. Repeat for other plates having an appropriate number of cells per plate. E.
As a control, try staining one of the plates with ampicillin. ampicillin. All the colonies colonies on this plate should have a halo.
VII.Calculating the Results A. Calculate the concentration of plasmid-containing cells (P) and plasmid-lacking cells (N) in the fermentation culture using the
colony-count data and the known dilution factors. Estimate the standard deviation of the plating technique. B.
Calculate the fraction of cells containing the plasmid at the beginning and end of the experiment using Equation 6.
C.
Calculate the the number of generations between the initial and final samples using Equation 5, and the probability of plasmid loss in each medium using Equation 4.
Figure 1. General Plasmid and Plasmid with DNA Insert
Immobilized Cell Biocatalysts Introduction Immobilization of cells and enzymes offers several advantages for bioreactor operation. Immobilization allows the biocatalysts to be retained in the reactor, thus conserving biocatalysts, allowing higher biocatalysts concentrations, and eliminating the need for downstream separation of the biocatalysts from the product. Immobilization can also increase the stability of the biocatalysts in some cases (Petersen, 1991). However, immobilization also has disadvantages. When the biocatalysts are entrapped within a porous matrix, the matrix provides an additional resistance to substrate mass transfer. The simultaneous diffusion and consumption of substrate within the catalyst particles cause the substrate concentration to decrease with depth into the matrix. matrix. The concentration often falls to zero inside the pa rticles, thus reducing the biocatalyst effectiveness. Under these conditions, substrate diffusion is the rate-limiting rate-limiting factor, and bioreactor design requires the ability to accurately predict the diffusivity of the substrate substrate through the catalyst matrix. The variable that characterizes the rate rate of substrate diffusion diffusion is the effective diffusivity. diffusivity. The purpose of this experiment is to demonstrate how an effective diffusivity can be measured measured using an unsteady-state approach. The diffusivity of calcium chloride (CaCl2) in a calcium-alginate gel matrix will be experimentally determined as a function of temperature and an d compared to literature values.
Background Techniques to immobilize biocatalysts can be classified into two groups: chemical and physical. In chemical immobilization, a covalent bond is formed between the solid phase and the biocatalyst. This type of immobilization is most frequently used used for enzymes or dead cells, because the harsh reaction conditions required are generally lethal to cells. Physical immobilization can be achieved in a variety of ways: entrapment within a porous matrix, adsorption, flocculation (i.e., self-aggregation), or containment behind a membrane. Polysaccharide gels, such as alginate and κ-carrageenan, are commonly used porous matrices for entrapment of living cells. Uniformly sized spheres containing viable viable cells are easily made under mild conditions conditions with these gels. The porosity, or void fraction, fraction, of these gels is quite high—typically greater than 95%--so that substrates and products can readily diffuse throughout the gel. Alginate, a skeletal component of seaweed, seaweed, is a watersoluble polysaccharide containing multiple carboxylic acid groups. Divalent cations, 2+ such as Ca , create ionic bonds that crosslink adjacent alginate molecules to form a solid gel. The pore size, molecular permeability, and degree of crosslinking of the gel vary with the molecular weight of the alginic acid. Spherical biocatalysts are formed formed by dripping an aqueous solution containing sodium alginate and the cells of interest into a hardening solution containing a calcium salt. After the drops have solidified, solidified, they are 2+ transferred into an appropriate growth growth medium. Molecules that can either scavenge Ca + ions, (e.g., citrate and phosphate) or displace them (e.g., Na ions) are used sparingly in the medium, because they can soften or dissolve the gel.
Experimental Equipment and Procedure The experimental apparatus used to produce the alginate alginate beads is shown in Figure 1. It consists of a magnetic stirrer, plastic tubing, a funnel, a plastic nozzle, and a c lamp for the tubing. The alginate solution drips into the the CaCl2 hardening solution, where the drops solidify into into spheres. About 50 mL mL of beads are needed for each run. run. However, by making at least 100 mL, one batch of beads can be used while the CaCl2 is being removed from the other batch. In this way, experimental data may be taken continuously. After the beads are incubated in the hardening solution, and after each experimental run, they should be incubated in several times their volume of RO water to remov e the unbound CaCl2. The RO water should be changed every 5-10 minutes minutes for about an hour. At least 25 beads should be measured to determine the average bead diameter. The experimental equipment used to measure the rate of diffusion of CaCl2 into the alginate beads is shown in Figure 2. It includes a water-bath shaker, shaker, a conductivity probe and meter, and the Opto-22 data-acquisition system. The voltage output from the conductivity meter must be calibrated. Standard solutions containing containing known concentrations of CaCl2 in RO water are warmed to the desired temperature in 250 mL Erlenmeyer flasks. The probe is placed in a flask, shaking is initiated, and the conductivity output from the meter is recorded. Because the conductivity is proportional to the concentration in the concentration range used, a two-point, linear calibration curve may be used. Further details of the calibration procedure are given in the Appendix. Because conductivity is a function of o f temperature, the calibration must be performed at the same temperature as the experimental runs. In addition, the beads, CaCl2 solution, and conductivity probe should all be in thermal equilibrium before the run is started. The experimental runs are initiated by dropping about 50 mL of alginate beads that are initially devoid of unbound CaCl2 into a 10 g/L CaCl2 solution. The conductivity probe is then quickly replaced into the the solution, and shaking is initiated. initiated. The conductivity is recorded by the Opto-22 system until steady state is reached. At that point, the next run run can be initiated with the second batch of beads, and the first batch can be incubated in RO water to prepare them for the following run. To get enough data to analyze the effects of temperature and perform replicates, several successful runs (about three three per lab period) are needed. However, each run requires 3040 minutes of data logging plus additional time for calibrating the probe, changing solutions, etc. Experience has shown that early preparation, good planning, and time management are particularly important in succeeding in this experiment.
Theoretical Analysis Radial diffusion of substrates through the highly porous matrix of alginate can be modeled using Fick’s law: ∂ C s J s = - D e ∂r
where
Js = the diffusive flux of the substrate De = the effective diffusion coefficient Cs = the concentration of the substrate in the sphere r = radial coordinate
Substituting Fick’s law into the unsteady-state mass-balance equation on substrate within a sphere, and assuming that convective flux inside the sphere is negligible, gives:
∂ C s 1 ∂ 2 ∂ C s = ( r D e ) ∂t r 2 ∂r ∂r
(2)
where t = time The experiment can be conveniently con veniently run in two different ways: (1) with CaCl2 diffusing into beads initially equilibrated with RO water and (2) with CaCl2 diffusing out of beads initially equilibrated with CaCl2 solution into RO water. The following initial and boundary conditions are appropriate when spheres initially devoid of solute are submerged in a well-mixed, solute-containing liquid phase o f constant volume, with no mass-transfer resistance between the bulk liquid and the beads:
at t = 0; 0 < r < ; C s = 0
(3)
at t = 0; r > R; C L = C Lo
(4)
at t > 0; r = 0 :
t > 0; r = R; V L
where
∂ C S =0 ∂r
∂ C L ∂ C s = K p A s D e | ∂t ∂r r = R
(5)
(6)
R = radius of the beads CL = concentration in the liquid phase CLo = initial concentration in the liquid phase VL = volume of the liquid phase K p = partition coefficient between liquid and bea ds As = surface area of the beads
Crank (1975) gives the following series solution for the liquid-phase solute c oncentration in terms of the diffusion coefficient, the ratio of the liquid volume to the bead volume (α), the radius of the beads, and several qn values:
C L =
α C Lo {1 + 1 + α
∞ ∑ n=1
2
- D q t 6(1 + α ) exp( e 2 n ) R } 2 2 9 + 9α + qn α
(7)
Values for the qn’s are given by the non-zero non -zero solutions of the following equation: tan( qn ) =
3 qn
(8)
3 + α qn
For the second case, where CaCl2 is diffusing into beads initially devoid of the salt, the solution is: 2
∞ 6 α (1 + α ) exp( - De qn t ) 2 C po R − ∑ = {1 } C L 2 1 + α 9 + 9α + q n α 2 n=1
(9)
Where C p0 = initial CaCl2 concentration inside the bead. A Fortran computer program has been written to automatically fit the model described above to the experimental data. The program uses a non-linear optimization optimization subroutine, Patern, to determine the values of De and α that minimize the deviation between the model predictions and the experimental data. Newton’s method is used used to find the roots (qn) of Equation 8. Further details about the program are given in the the Appendix. A copy of the program will be transferred to your 3 ½ in floppy disk for you to use in analyzing your data. Published values for the aqueous a queous diffusivity of CaCl2 may be found in th handbooks of physical properties, such as the 57 Edition of the CRC Handbook of Chemistry and Physics , for comparison with your experimental values. Accurate determination of the effective diffusion coefficient requires mass transfer resistances other than that of the catalyst matrix be made negligible. The stagnant fluid film surrounding a submerged object can, under certain conditions, create a significant mass transfer resistance. resistance. The thickness of this film film (δ), and hence its resistance, is inversely proportional to the interphase mass-transfer coefficient, as shown below (Nguyen, 1986): δ =
Dm k
(9)
where Dm is the diffusivity of CaCl2 into free water. The interphase mass-transfer mass-transfer coefficient (k) is defined by the following equation: N s = k( C L - K p C S )
where Ns is the convective flux from the bulk fluid into the sphere.
(10)
Experimental data giving k as a function of the fluid velocity, and the geometry and physical properties of the system are typically correlated in dimensionless form, as shown below: where
Sh = f(Re,Sc) Sh = Sherwood number = kd/Dm Sc = Schmidt number = µ/ρDm Re = Reynolds number = ρud/µ d = characteristic length (e.g., bead diameter) Dm = diffusion coefficient of the substrate in the fluid pha se µ = fluid viscosity ρ = fluid density u = velocity of fluid relative to the sphere
(11)
In general, Sh increases with Re to about the 0.6 to 0.8 power. Thus, the mass-transfer resistance may be minimized (i.e., k may be maximized) by maximizing the fluid velocity v elocity relative to the catalyst particles. particles. Experimentally, this result is achieved by sufficiently agitating the fluid. Thus, the experiment should be performed at a high agitation rate (about 300 rpm). The effect of temperature on the diffusivity can be predicted from one of the commonly used correlations used to predict diffusivities, such as the Wilke-Chang correlation.
Symbols
α = ratio of liquid volume to bead volume δ = film thickness µ = fluid viscosity ρ = fluid density As = surface area of the beads Ci = the concentration of component i CL = concentration in the liquid phase CLo = initial concentration in the liquid phase Cs = solute concentration in the beads d = bead diameter De = effective diffusion coefficient Dm = diffusivity into water Ji = the diffusive flux of component i k = external mass transfer coefficient K p = partition coefficient between liquid and solid phase (beads) n = eigenvalue index Ni = total flux of component i qn = eigenvalues of the solution for CL/CLo r = radial coordinate R = radius of the beads r = radial coordinate Re = Reynolds number = ρud/µ Sc = Schmidt number = µ/ρDm
Sh = Sherwood number = kd/Dm t = time u = liquid velocity relative to beads VL = volume of the liquid phase
References Crank, J., Mathematics J., Mathematics in Diffusion, Diffusion, Claredon Press, Oxford, 1975, pp. 56-60, 93-97. Cussler, E. L., Diffusion: L., Diffusion: Mass Transfer in Fluid Systems, Systems, Cambridge University Press, New York, 1984, pp.19-21. Freund, Vicki, “Using DIFF to Determine Diffusion Coefficients”, Department of Chemical Engineering, Michigan State University, photocopy Itamunoala, G. F., “Effective Diffusion Coefficients in Calcium Alginate Gel”, Biotechnology Progress, Progress, 3, June 1987, pp. 115-121. Kennedy, John F. and Cabral, Joaquim M. S., “Immobilized Living Cells and Their Applications” Applied Applications” Applied Biochemistry and Bioengineering Volume 4: Immobilized Microbial Cells, Cells, Academic Press, New York, 1983. Nguyen, An-Lac, and J. H. T. Luong, “Diffusion in κ-carrageenan Gel Beads” Biotechnology and Bioengineering , vol. 28,pp. 1261-1267, J. Wiley and Sons, 1986 Petersen, James N., Davidson, Brian H., and Charles D. Scott, “Minimizing the Errors Associated with the Determination of Effective Diffusion Coefficients when using Spherical Cell Immobilization Matrices” Biotechnology Matrices” Biotechnology and Bioengineering , vol 37, pp. 386-388, John Wiley and Sons, Inc., 1991.
Appendix 1: Experimental Supplies Supplies and Equipment Equipment sodium alginate calcium chloride graduated cylinders funnel rubber tubing screw clamps conductivity meter Reverse osmosis (RO) water balance
Appendix 2: Details of of Experimental Procedure I
Preparation of Alginate Solution (must be done prior to the day beads are to be made) A. A 2% alginate stock solution should be available in the refrigerator for your use. If there is less than about 200 mL remaining after you have prepared your beads, please make more for the next group to use. The solution is prepared as described below. The key to successfully successfully making the solution is avoiding the formation of large clumps as the alginate powder is being added to the water.
II
B.
Add 500 to 1000 mL of warm (about 50°C) RO water to an Erlenmeyer flask, along with a large stirring stirring bar. Set the magnetic stirring stirring plate to give a strong strong vortex. Then, sprinkle the alginic acid or sodium alginate powder onto the surface of the water so that the vortex rapidly disperses the powder throughout the solution as small clumps. The solution thickens considerably as the alginate dissolves, so try to disperse the alginic acid as quickly as possible without making large clumps. Reduce the agitation speed so that it no longer entrains air, and allow the remaining clumps c lumps to dissolve. Occasionally autoclaving may be necessary to complete the dissolving process.
C.
Once the alginate solution solution has completely dissolved, dissolved, allow allow the solution to stand so that entrapped air bubbles may escape. Store the alginate solution in the the refrigerator.
Preparation of Alginate Spheres A. Set up the apparatus shown in Figure 1. The plastic nozzle should have a tip diameter between 2 and 5 mm. Locate the funnel as high as possible possible to give time for the drops to acquire a spherical shape before entering the hardening solution. B.
Prepare about 300 mL of 15 g/L, aqueous, CaCl2 hardening solution in a 600 mL beaker. Position the beaker so the drops fall closer to the wall of the beaker than the center. The magnetic stirring rate rate should be just adequate to carry each drop away from the dripping zone before the next one lands. Too much agitation causes the drops to be deformed.
C.
Slowly open the the clamp to allow the the alginate solution solution to drip at a rate of about 2 drops per sec. When the desired volume of beads has been produced, close the clamp, and let the beads cure in the hardening solution for 30 to 60 minutes. The alginate spheres shrink during curing; a bout 200 mL of alginate solution is needed to make 100 mL of beads.
D. Once the beads have cured, they should be incubated in a large volume of RO water to remove excess CaCl2. The experiments are generally started started with beads that are devoid of CaCl2. About 30 minutes is required for for the excess salt
to be removed. The RO water should be changed during this time to maintain a low salt concentration surrounding the beads. III Calibration of the Conductivity Meter A. While the beads are being made, the shaker shaker should be warmed up to to the desired desired temperature. (This process can be sped up by replacing replacing some of the water with with RO water that has been heated in the microwave oven). The experimental temperature should be at least 15 degrees above room temperature to prevent a temperature overshoot due to the mechanical agitation. B.
Using RO water, prepare several CaCl2 standard solutions in the range of 0 to 10 g/L CaCl2. Place about 150 mL of each standard in in a labeled, 250 mL mL Erlenmeyer flask, and cap with either either Parafilm or a rubber rubber stopper. Warm the standards to the desired temperature in the incubator.
C.
Rinse the conductivity probe in RO water, and place it in the 0% CaCl2 standard. The probe-stopper assembly should fit fit snugly into the neck of the flask so the hole in the the probe is well below the the liquid level. Also, the cord between the conductivity probe and meter should be securely taped down to minimize noise in the voltage signal.
D. Set the function dial on the conductivity meter to “conductance”, and the range dial to “20 mmho”. mmho”. Note: Electrical conductance is the inverse of resistance; the unit of conductivity, the mho, is given the reverse spelling of the ohm, and the symbol of the mho is the symbol for the ohm (Ω) turned upside down. IV. Conducting the Experimental Runs Note: It is most efficient to use two sets sets of beads. At any time, one set would be used for one diffusion experiment while the other set would be equilibrating in a standard solution (either RO water or a 10g/l solution of CaCl2). In this way, at the end of each run, another set of beads is ready to be used for the next run. A. Switch on the constant-temperature constant-temperature water bath, and set it to the temperature at which the calibration was performed. The warm-up process can be sped up by replacing some of the water with RO water that has been heated in the microwave oven. B.
Warm the conductivity probe, alginate beads and CaCl2 solution to be used for the experimental run up to the desired desired temperature. Set the function dial on the conductivity meter to “conductance”, and the range dial to “20 mmho”. Make sure that the reading on the conductivity meter is not off the scale. If it is, adjust the dial setting to bring the conductivity back onto the scale.
C.
Once the standards have achieved the setpoint setpoint temperature, measure their conductivity to confirm that the calibration curve has not changed.
D. Determine the volume of each set of beads: 1. Pour between 100 and 130 mL of RO water into a 250 mL graduated cylinder, and record the volume. 2. Collect about 50 mL of beads and spread them out onto a damp paper towel to remove excess water. 3. Transfer the beads into the graduated cylinder. Record the final volume, and determine the volume of the beads by difference. F.
Collect the beads, and remove excess water as before. Transfer the beads to a dry, 250 mL Erlenmeyer flask. Cap the flask, and place it in the incubator to keep the beads at the desired temperature.
G. Transfer a measured volume (about 150 mL) of 10 g/l CaCl2 solution into a 250 mL Erlenmeyer flask, and securely fasten the conductivity probe-stopper assembly into the neck of the flask. Mount the flask into the water bath, and allow the temperature to equilibrate with the water bath. H. Set the shaker at the speed used for the calibration calibration runs. I.
When ”Start” is pressed on the Opto 22 screen, the system saves data to the the end of a data file that can be opened and manipulated into Excel as a spreadsheet. The Opto 22 display screen contains information about the location of the folder in which the file is stored. The file will contain data from previous groups’ runs if it has not been erased before the current group begins to use it. To calibrate the voltage readings, the display must be in “Start” mode.
D. Monitor the experiment using the Opto 22 display screen. V. Data Recovery and Analysis A. Copy the Opto 22 version of the data to a floppy disk. B.
Import the data file into Excel for for manipulation. Convert the time data from from seconds to minutes to be consistent with the Fortran program “DIFFTR7A”. Create a new file containing only the time data (in minutes) in the left-hand column and the concentration data (in g/L) in in the right-hand column. The time data should start at a value of zero minutes in the upper left-hand corner of the data file. If data were recorded prior to the addition of the beads to the CaCl2 solution, they should be deleted. Count the number of time points in the file; the Fortran file will ask for this value.
C.
Run DIFFTR7A.EXE DIFFTR7A.EXE using the command command “a:difftra7”, “a:difftra7”, if the program is on a floppy disk.
D. The computer will then ask whether you wish to solve Case 1 or 2. Enter 1 if the CaCl2 was diffusing into the beads and 2 if the CaCl2 was diffusing out of the beads..
E.
The program will request request the name of the file containing the data and the number of data points. Enter path and filename in single single quotes, followed by a comma and the number of time points (e.g., ‘a:\filename.csv’,90)
F.
The number of qn values to use in the series solution should then be input. Six is usually sufficient, although more may be used.
A. The average radius of the beads (not the diameter) should then be input in in units of cm. The program will determine the values of α and De that minimize the error between the model model prediction and experimental data. Information will be shown on the screen tracking the progress of the program.
B.
Experience has shown that that inputting inputting the initial concentration evaluated from the data gives better results than inputting the target value (e.g., (e.g., 10.0 g/L). Some water can be carried over with the beads that may cause the initial concentration to differ slightly from 10 g/L. To obtain the initial value, plot the experimental data as concentration vs. time. Draw a smooth curve through the data back to t=0. Use the value the smooth curve has at t=0 as the initial concentration.
C.
When the the program stops, record the “cost”, “parameter “parameter 1”, and “parameter 2”. 2 These values correspond to the error of the model fit, De in units of cm /min, and α. Also, record the listed values of qn.
D. To evaluate the fit fit of the model, model, use the qn, , and De values to calculate the theoretical curve using the series solution, and plot it on the same graph as the experimental data. Also, calculate an α value from the measured volumes of the solid and liquid phases, and compare the two values. Appendix 3: Difftr7a Program to Evaluate D and
C SUBROUTINE PROC FOR OPTIMIZATION SUBROUTINE PATERN C C PURPOSE: TO ESTIMATE DIFFUSION COEFFICIENTS BY READING IN DATA C CONTAINING TIME -VS- CONCENTRATION VALUES. C C C C C
MAIN PROGRAM PROGRAM CHE460 IMPLICIT DOUBLE PRECISION(A-H,O-Z) INTEGER NDATA,NP,NPASS,IO, NED COMMON C(4000), T(4000), Q(10000), NDATA, X, NQ, R COMMON NTYPE, XB DIMENSION P(10), STEP(10)
CHARACTER CONCEN*20 C C
C C C 5
SEARCH INITIALIZATION P(1)=5.0E-4 P(2)=4.00 STEP(1)=1.0E-5 STEP(2)=0.1 NP=2 NPASS=3 IO=3 READ IN DATA WRITE(*,*)’WHAT TYPE OF DIFFUSION PROBLEM DO YOU WISH TO’ WRITE(*,*)’SOLVE? ENTER THE NUMBER NUMBER FOR THE DESIRED DESIRED TYPE.’ WRITE(*,*) WRITE(*,*)’1. DIFFUSION FROM LIQUID INTO INITIALLY’ WRITE(*,*)’ SOLUTE FREE SPHERES.’ WRITE(*,*)’2. DIFFUSION FROM SPHERES INTO INITIALLY’ WRITE(*,*)’ SOLUTE FREE LIQUID.’ READ(*,*) NTYPE WRITE(*,*)
C IF(NTYPE.EQ.1) THEN WRITE(*,*)’ENTER INITIAL LIQUID CONCENTRATION IN g/L’ READ(*,*) X WRITE(*,*) ELSEIF(NTYPE.EQ.2) THEN WRITE(*,*)’ENTER INITIAL BEAD CONCENTRATION IN g/L’ READ(*,*) XB WRITE(*,*) ELSE WRITE(*,*)’YOU DID NOT ENTER A 1 OR A 2. TRY AGAIN’ WRITE(*,*) GOTO 5 ENDIF C WRITE(*,*)’ENTER THE NAME OF THE TIME-CONCENTRATION FILE’ WRITE(*,*)’AND THE NUMBER OF DATA POINTS’ READ(*,*) CONCEN, NDATA WRITE(*,*) C OPEN(UNIT=20,FILE=CONCEN,STATUS=’OLD’) C C WRITE(*,*)’ENTER THE NUMBER OF Q VALUES DESIRED’ READ(*,*) NQ
WRITE(*,*) WRITE(*,*)’ENTER THE RADIUS OF THE BEADS’ READ(*,*) R WRITE(*,*) C
350 150 C C C C
WRITE(*,*) WRITE(*,*)’CONCENTRATION(g/L) DO 150 J=1,NDATA READ(20,*)t(J),c(J) WRITE(*,350)C(J),T(J) FORMAT(6X,F6.2,18X,F6.2)
TIME(MIN)’
CONTINUE
START SEARCH CALL PATERN(NP,P,STEP,NPASS,IO,COST)
C C C C300
300
346
C C C C C C C C C C C C C C
SEARCH COMPLETE, PRINT RESULTS PRINT 300, P(1), P(2), COST FORMAT(F10.3,10X,F10.3) OPEN (UNIT=30,FILE=’FOR30.DAT’,STATUS=’NEW’) WRITE (30,300) P(1),COST FORMAT(E10.3,5X,F6.3,5X,E10.3) DO 346 L=2,NQ+1 WRITE (*,*)Q(L) CONTINUE STOP END
THIS FILE IS A PAIR OF SUBROUTINES WRITTEN TO BE COMPATIBLE WITH THE OPTIMIZATION SUBROUTINE PATERN. THEY SIMULATE A PROCESS USING DISCRETE DIFFERENCE EQUATIONS AND COMPARE THE SIMULATION OUTPUT WITH THE ACTUAL OUTPUT (READ IN THROUGH A DATA FILE), CALCULATING AN ERROR OR “COST” ASSOCIATED WITH THAT SIMULATION. PATERN USES THESE SUBROUTINES ITERATIVELY IN ORDER TO FIND THE OPTIMUM SET OF TRANSFER FUNCTION PARAMETERS TO FIT THE DATA.
SUBROUTINE PROC(P,COST) IMPLICIT DOUBLE PRECISION (A-H,O-Z) COMMON C(4000), T(4000), Q(10000), NDATA, X, NQ, R
COMMON NTYPE, XB DIMENSION P(10), STEP(10), CONC(100) C C C
INITIALIZE ARRAYS AND DEFINE PARAMETERS D=P(1) ALPHA = P(2) ERROR=0.0 TOTAL=0.0
C C PURPOSE: TO CALCULATE THE Q VALUES BASED ON THE ALPHA GIVEN C C Program solver *Solves for the first N solutions of the variable QN given alpha *USING NEWTON’S METHOD *DEFINE THE FUNCTION F WHICH MUST = 0 TO OBTAIN A QN VALUE *FP = THE DERIVATIVE OF THE FUNCTION F C C *TOL1 IS THE CONVERGENCE CRITERION TOL1 = .00000001 QNOLD IS THE CURRENT VALUE OF QN QNOLD = 3 NC = 1 *ALL OF THE IF STATEMENTS BELOW INVOLVING ALPHA GIVE THE COMPUTER *A FIRST GUESS CLOSE TO THE TABULATED VALUES FROM CRANK IF (ALPHA .LT. .6667) THEN QNOLD = 4 ENDIF IF (ALPHA .LT. .4286) THEN QNOLD = 4.18 ENDIF *THIS IS THE MASTER LOOP IT IS CONTROLLED BY THE NUMBER OF QN VALUES DESIRED 110 IF (NC .LE. NQ) THEN FPOLD = 1/((DCOS(QNOLD))**2)-(3*ALPHA*QNOLD**2 + -6*ALPHA*QNOLD+9)/((3+ALPHA*QNOLD**2)**2) FVAL = DTAN(QNOLD)- 3*QNOLD/(3+ALPHA*QNOLD**2) IF (FPOLD .EQ. 0) THEN WRITE (*,*) ‘NEWTONS METHOD FAILS’ NC=NQ+1 ELSE QNNEW=QNOLD-(FVAL/FPOLD) QNOLD=QNNEW NED=NED+1 ENDIF
C C120 C C
IF (ABS(FVAL) .LT. TOL1) THEN NC = NC + 1 QNOLD=QNOLD+2 WRITE (*,120) ‘QN’,’F(QN)’,’FP(QN)’ FORMAT (15X,A6,15X,A6,25X,A6) WRITE (*,*) QNNEW,FVAL,FPOLD
Q(NC) = QNNEW WRITE (*,*) NC, Q(NC),NED,ALPHA NED = 0 C ENDIF GOTO 110 ENDIF C C C C
10
20
40
30 300
CALCULATE COST IF(NTYPE.EQ.1) THEN DO 20 J=1,NDATA DO 10 I=2,NQ+1 TERM1=(6*(1+ALPHA)*EXP(-D*Q(I)**2*T(J)/R**2)) TERM=TERM1/(9+9*ALPHA+Q(I)**2*ALPHA**2) TOTAL=TOTAL+TERM CONTINUE CONC(J)=((X*ALPHA)/(1+ALPHA))*(1+TOTAL) ERR=ABS(C(J)-CONC(J)) ERROR=ERROR+ERR COST=ERROR TOTAL=0.0 CONTINUE GOTO 300 ELSE DO 30 J=1,NDATA DO 40 I=2,NQ+1 TERM1=(6*(1+ALPHA)*EXP(-D*Q(I)**2*T(J)/R**2)) TERM=TERM1/(9+9*ALPHA+Q(I)**2*ALPHA**2) TOTAL=TOTAL+TERM CONTINUE CONC(J)=(XB/(1+ALPHA))*(1-TOTAL) ERR=ABS(C(J)-CONC(J)) ERROR=ERROR+ERR COST=ERROR TOTAL=0.0 CONTINUE ENDIF RETURN
END C C C C SUBROUTINE BOUNDS(P,IOUT) IMPLICIT DOUBLE PRECISION (A-H,O-Z) COMMON C(4000),T(4000),Q(10),NDATA, X, NQ, R COMMON NTYPE, XB DIMENSION P(1), STEP(1) IOUT=0 IF(P(1).LT.0.OR.P(2).LT.0) IOUT=1 RETURN END C SUBROUTINE PATERN(NP,P,STEP,NRD,IO,COST) C--------THE SIZE OF B1,B2,T,AND S NEED ONLY BE EQUAL TO THE NUMBER OF PAR IMPLICIT DOUBLE PRECISION (A-H,O-Z) DIMENSION P(1000),STEP(1000),B1(100),B2(100),T(100),S(100) C----------------------------------------------------------C THE FOLLOWING COMMAND ALLOWS PATERN TO USE AN INTEGER C VARIABLE AS THE THIRD PARAMETER P(3). C NSRC =3 P(NSRC) = IDINT(P(NSRC)) C------------------------------------------------------------C-----STARTING POINT L=1 ICK=2 ITTER=0 DO5 I=1,NP B1(I)=P(I) B2(I)=P(I) T(I)=P(I) 5 S(I)=STEP(I)*10. C-----INITIAL BOUNDARY CHECK AND COST EVALUATION CALL BOUNDS(P,IOUT) IF(IOUT.LE.0)GOTO10 IF(IO.LE.0)GOTO6 WRITE(*,1005) WRITE(*,1000)(J,P(J),J=1,NP) 6 RETURN 10 CALL PROC(P,C1) IF(IO.LE.0)GOTO11 WRITE(*,1001)ITTER,C1 WRITE(*,1000)(J,P(J),J=1,NP)
C-------BEGINNING C-------BEGINNING OF PATTERN SEARCH STRATEGY 11 DO99 INRD=1,NRD DO12 I=1,NP 12 S(I)=S(I)/10. C-------------------------------------------------------------S(NSRC) = 1.0001 C-------------------------------------------------------------IF(IO.LE.0)GOTO20 WRITE(*,1003) WRITE(*,1000)(J,S(J),J=1,NP) 20 IFAIL=0.0 C-----PRETURBATION ABOUT T DO30 I=1,NP IC=0 21 P(I)=T(I)+S(I) IC=IC+1 CALL BOUNDS(P,IOUT) IF(IOUT.GT.0)GOTO23 CALL PROC(P,C2) L=L+1 IF(IO.LT.3)GOTO22 WRITE(*,1002)L,C2 WRITE(*,1000)(J,P(J),J=1,NP) 22 IF(C1-C2)23,23,25 23 IF(IC.GE.2)GOTO24 S(I)=-S(I) GOTO21 24 IFAIL=IFAIL+1 P(I)=T(I) GOTO30 25 T(I)=P(I) C1=C2 30 CONTINUE IF(IFAIL.LT.NP)GOTO35 IF(ICK.EQ.2)GOTO90 IF(ICK.EQ.1)GOTO35 CALL PROC(T,C2) L=L+1 IF(IO.LT.2)GOTO31 WRITE(*,1002)L,C2 WRITE(*,1000)(J,T(J),J=1,NP) 31IF(C1-C2)32,34,34 32 ICK=1 DO33 I=1,NP B1(I)=B2(I) P(I)=B2(I) 33T(I)=B2(I)
GOTO20 34 35
C1=C2 IB1=0 DO39 I=1,NP B2(I)=T(I) IF(ABS(B1(I)-B2(I)).LT.1.0E-20)IB1=IB1+1 39 CONTINUE IF(IB1.EQ.NP)GOTO90 ICK=0 ITTER=ITTER+1 IF(IO.LT.2)GOTO40 WRITE(*,1001)ITTER,C1 WRITE(*,1000)(J,T(J),J=1,NP) C-----ACCELERATION STEP 40 SJ=1.0 DO45 II=1,11 DO42 I=1,NP T(I)=B2(I)+SJ*(B2(I)-B1(I)) C-----------------------------------------------------------------IF(I.EQ.NSRC)T(I)=IDINT(T(I)) C---------------------------------------------------------------42 P(I)=T(I) SJ=SJ-.1 CALL BOUNDS(T,IOUT) IF(IOUT.LT.1)GOTO46 IF(II.EQ.11)ICK=1 45 CONTINUE 46 DO47 I=1,NP 47 B1(I)=B2(I) GOTO20 90 DO91 I=1,NP 91 T(I)=B2(I) 99 CONTINUE DO100 I=1,NP 100 P(I)=T(I) COST=C1 IF(IO.LE.0)RETURN WRITE(*,1004)L,C1 WRITE(*,1000)(J,P(J),J=1,NP) RETURN 1000 FORMAT(3(35X,I7,5X,E13.6/)) 1001 FORMAT(//1X13HITERATION NO. ,I5/5X,5HCOST= ,E15.6,20X, 1 10HPARAMETERS) 1002 FORMAT(10X3HNO.,I4, 8X5HCOST=,E15.6) 1003 FORMAT(/1X28HSTEP SIZE FOR EACH PARAMETER )
1004 FORMAT(1H113HANSWERS AFTER ,I3,2X,23HFUNCTIONAL EVALUATIONS // 1 5X5HCOST=,E15.6,20X,18HOPTIMAL PARAMETERS ) 1005 FORMAT(1H135HINITIAL PARAMETERS OUT OF BOUNDS END
)
Figure 1. Apparatus for Forming Gel Beads
Figure 2. Experimental Setup for Determination of Effective Diffusion Coefficients
Mass Transfer Introduction Fermentation can be loosely defined as the production of chemicals using microorganisms as catalysts catalysts for the reaction. Fermentation is used to produce produce a variety of commercial products including alcohols, organic acids, and pharmaceuticals. Many fermentations are carried out using aerobic microorganisms that consume dissolved oxygen from the fermentation broth. Oxygen is replenished in the liquid phase by sparging air or pure oxygen through the the fermentation broth. Because oxygen transfer is often rate-limiting in aerobic fermentations, the ability to accurately measure and predict the oxygen mass-transfer rate is needed for bioreactor design and optimization (Roberts et al., 1992). The variable used to characterize the mass-transfer rate rate is the volumetric mass transfer coefficient (k La). The objective of this experiment is to determine k La for oxygen transfer in a stirred-tank stirred-tank fermenter using the “static-gassing-out” method. The k La data will be correlated as a function of superficial gas velocity, coalescence properties of the liquid, impeller rate and impeller type. The results will be compared to published results.
Background The productivity of aerobic fermentations is often limited by the concentration of oxygen in the fermentation broth (C). This concentration is determined by the balance between the rate of oxygen consumption by the cells and the rate of oxygen transfer into the liquid from the gas. The volumetric oxygen transfer rate is given by the product of k La and the * * driving force for mass transfer, C - C, where C is the liquid-phase oxygen concentration that would be in equilibrium with the gas phase (i.e., the the oxygen solubility). The driving force is inherently low, due to low aqueous solubility of oxygen. The liquid-phase oxygen concentration in equilibrium with air under ambient conditions is only about 10 parts per million (ppm). By contrast, a typical aerobic yeast fermentation requires over 750 times that amount (Atkinson, 1983). Efforts to increase the oxygen mass-transfer rate typically focus on increasing k La. Two terms make up k La: k L, the liquid-film mass-transfer coefficient, and a, the interfacial area per unit liquid volume. Although k L is affected by surface-active substances in the medium that foul the interface between the bubble and liquid, it is difficult to control co ntrol this effect. Thus, k La is generally increased by increasing a. Because the surface area of spheres is proportional to the diameter squared, while the volume is proportional to the diameter cubed, a is inversely proportional to the bubble diameter. Consequently decreasing the bubble diameter is an effective strategy for increasing k La. After the gas is injected through the sparger into the liquid phase, the average bubble size may change due to bubble coalescence (merging of two bubbles) and bubble disintegration (bubble breakup). breakup). The equilibrium bubble size size is determined determined by the balance between hydrodynamic forces that tend to deform the bubble, leading to bubble breakup, and surface tension, which tends to hold the bubble in a spherical shape. Solutes that accumulate at the gas-liquid interface (e.g., surfactants) strongly influence
the surface tension, and hence k La. Surfactants that are either initially present in fermentation medium or produced during the fermentation may cause a foaming problem. Foaming is combated by the addition of chemical antifoams. Not surprisingly, both the surfactants and antifoams have a strong effect on the k La value. Thus, it is important to measure k La under conditions as similar as possible to those to be used in industrial practice. The most common methods to increase a in stirred tanks are to increase the agitation rate and to increase the gas sparging rate. Increasing the impeller rate increases increases impeller shear, thus increasing bubble breakup and increasing a. Smaller bubbles also have a smaller rise velocity, and consequently a greater residence time in the liquid. The longer the bubbles remain in the reactor, reactor, the more oxygen may be transferred. transferred. Increasing the sparging rate increases the number of bubbles present in the liquid. The volumetric mass-transfer coefficient is a function of the tank a nd impeller geometry, the agitation power, the rate and method of sparging, and the physical properties of the liquid. The geometry of the mixing tank, power input and the impeller style style are specified during the design of the system. system. The tank geometry may be characterized by several geometric ratios known as “shape “shape factors” (McCabe, Smith, and Harriot, 1985). Mixing vessels are often designed with typical shape factors, for which experimental data exist. The type of impeller used for agitation strongly affects both the mass-transfer coefficient and power requirement. Impellers are usually classified classified as either axial or radial flow. Axial-flow impellers displace the fluid downward along the axis of rotation, and radialflow impellers push the the fluid outward in the radial direction. Radial-flow impellers generate higher shear rates for better gas dispersion, but they consume more power than axial-flow impellers (Oldshue, 1990). Traditionally, commercial fermentors fermentors have used radial-flow impellers because of their excellent mass-transfer mass-transfer capabilities. However, the high operating cost of these impellers has led to the development of advanced, axial-flow impellers that promote mass transfer while still requiring less power than radial-flow impellers. This new generation of impellers has been developed using complex hydrodynamic modeling and laser-doppler anemometry (Oldshue, 1990).
Experimental Equipment and Procedure The equipment used for this experiment e xperiment includes two bioreactors having polarographic electrodes and amplifiers that that are connected to a Opto 22 data acquisition acquisition system. One of the bioreactors is a 1-L New Brunswick MultiGen fermentor, which has two 6-cmdiameter, 6-bladed, Rushton, radial-flow radial-flow impellers. Its glass tank has a 4.5-inch inner diameter and contains two 0.5-inch-wide 0.5-inch-wide baffles. The other bioreactor is a 10-L pilotscale fermenter. Figure 1 illustrates illustrates the experimental setup. The O2 probe and amplifier must be calibrated at the experimental temperature. temperature. The Multigen has an integrated temperature control system that ha s a thermistor probe and a resistance heating rod. The zero-oxygen output of the probe is first first recorded after sparging the water with N2 until the probe’s signal stabilizes. The amplifier’s output corresponding to the probe being in oxygen-free water is then recorded into Opto 22. Next, the water is sparged with air until the oxygen reading comes to steady state. The oxygen amplifier should then be adjusted to read 21 (the percentage of oxygen in air) by
turning the screw on the front front of the unit. The amplifier’s output corresponding corresponding to the probe being in water saturated with air is then recorded into Opto 22. From these two points, Opto 22 generates a calibration equation that automatically converts the mA signal from the amplifier amplifier into % saturation saturation units. Additional details of the calibration procedure are given in Appendix 2. An experimental run is carried out by making a step change either from air to N 2 or viceversa, and then recording the C data until until about 50% saturation is is achieved. This process generates enough data from one k La determination. The gas flow rate rate and impeller rate should then be increased to quickly bring the liquid to within about 5% of the equilibrium value. The process is then repeated using the other gas. In this way, two k La values are determined per cycle—one with air sparging, a nd the other with N2 sparging. This alternating process should be repeated over ove r a range of rpm values and gas flow rates. The data can be plotted as suggested by the integrated forms of the mass-balance equation, and k La may be determined from the slope of the linear portion of the curve using linear regression.
Theoretical Analysis The aqueous solubility of oxygen is controlled by the partial pressure of oxygen, the temperature and the presence of other solutes. The solubility of sparingly sparingly soluble gases such as oxygen are often modeled by Henry’s law: C = where
pG H
(1)
C = liquid phase concentration pG = the partial pressure of the gas H = Henry’s law constant
The static gassing out method is used to determine k La in this experiment. This method is based on the following unsteady-state mass balance on oxygen in the liquid phase of a stirred-tank reactor: d(CV) dt where
= k L a ( C * - C)V - (-r)V
k L = mass-transfer coefficient (length/time) a = specific interfacial area (area/volume) * C = liquid-phase oxygen concentration in equilibrium with the gas composition (mass/volume) C = liquid-phase oxygen concentration (mass/volume) t = time V = liquid volume in the reactor -r O2 con sumption (mass/volume⋅time) O2 = rate of oxygen consumption
When there is no oxygen-consuming reaction, r O2 O2 equals zero, and the mass balance becomes:
(2)
dC
= k L a ( C * - C)
dt * The method entails making a step change in C , and then monitoring the resulting change in C as a function of time using a dissolved-oxygen electrode. The C data are then graphically analyzed to obtain k La. The equation above may be integrated from when the step change is made (t = 0) to give C ln(1 - * ) = - ( k L a) t C
(3)
(4)
This equation is convenient when air or pure oxygen is the sparged gas. However, if * nitrogen (N2) or some other gas devoid of oxygen is sparged, then C equals zero. The unsteady-state equation may be divided by the liquid-phase oxygen concentration in * equilibrium with air (C air ) and integrated to give C ln( * ) = ( k L a) t C air
(5)
Note that in both of the integrated equations, the ratio of the actual dissolved oxygen concentration to that in equilibrium with air may be thought of as the fractional degree de gree of oxygen saturation, which is the experimentally measured variable. The data can be plotted in semi-log format, as suggested by Equations 4 and 5. The slope (k La) may be determined from the slope of the linear portion of the plot using linear regression. The experiments are quick to perform, so many runs can be made. Please do not include all the semi-log graphs in the report. One example plot for air sparging and one for nitrogen sparging would be sufficient to show the degree of linearity obtained in the experiments. If replicates are done, statistics may be p erformed, and the measured k La values can be plotted (with error bars) as a function of rpm and/or gas flow rate to develop a correlation. The unsteady-state method for determining k La is based on the assumption that the probe’s response accurately represents the liquid-phase oxygen concentration throughout the experiment. However, there is a time lag associated with the probe’s response. response. If the probe cannot respond rapidly enough to follow the true rate of change in the oxygen concentration, the resulting data will will be erroneous. The probe’s response time can be estimated by quickly transferring the probe from a solution having one oxygen concentration into a second (well-mixed) solution having a different oxygen concentration. A first-order model model can be fit to the data to determine the probe’s response-time constant. This constant should be considerably less than the largest k La values being measured to ensure accurate ac curate results. For common impeller and tank geometries, the mass-transfer coefficient is frequently correlated in terms of superficial gas velocity and powe r input as shown below: β
P γ k L a = α ( ) u s V
(6)
where
α, β, γ = constants dependent on hydrodynamic properties and the vessel geometry P/V = agitator power per unit reactor volume us = superficial gas velocity
The superficial gas velocity is the volumetric gas flow rate divided by the cross-sectional area of the tank. Correlations of this form are available in most Biochemical Engineering texts, including that by Bailey and Ollis (1986). An important result of this experiment will be experimental values of α, β, and γ. These values should then be compared to literature values from published correlations. More elaborate, dimensionless correlations are also available that consider the geometry g eometry of the system, and the physical properties of the fluid. The power input is a function function of the impeller rate rate (N). The relationship between power and impeller rate is often expressed in dimensionless terms by correlating the Power number (Pn) as a function of the Reynolds Number Number and Froude Number, which are defined below: P n =
Re = where
P g c ρ l N i3 Di5 ρ l Di2 N i µ
P = power output of mixer gc = Newton’s Law unit conversion constant ρl = liquid density Ni = impeller rate Di = impeller diameter µ = liquid viscosity
In turbulent flow, the Power number is essentially constant. constant. Under these conditions, the power input, and hence power-to-volume ratio, should increase with the impeller rate to the third power. Thus, a first approximation, the coefficient β may be estimated as the 3 slope of a logarithmic plot of k La vs. N , for a constant value of us. Similarly, γ may be estimated as the slope of a logarithmic plot of k La vs. us for a constant value of N. The values of β and γ determined using this approach may be compared to literature values. However, the α values should not be compared, because the differences in the measured variables would show up in the α value. Values of β of β and γ reported in the literature are average values obtained in different size reactors and under different conditions than those obtained in our laboratory. Thus when making a comparison, the literature values should not be considered “true” values from which to calculate a percent p ercent error, but merely another set of results with which to compare those obtained in your experiment. Correlations having the form of Equation 6 are available in the text for both bubble coalescing and bubble non-coalescing liquids. The experimental values of the constants con stants vary with the bubble-coalescence properties of
(7)
(8)
the liquid medium. A typical bubble coalescing liquid is distilled water, while a typical non-coalescing liquid is 0.1 M CaCl2 in tap water.
Symbols
α, β, α’, β’, γ = constants dependent on hydrodynamic properties and the vessel a = specific interfacial area (area/liquid volume) C = liquid phase oxygen concentration (mass/volume) * C = liquid phase oxygen concentration in equilibrium with the gas composition (mass/volume) H = Henry’s law constant k L = liquid phase mass-transfer coefficient (length/time) N = impeller rate , revolutions per minute pG = the partial pressure of the gas P/V = agitator power per reactor volume Qair = volumetric flow rate of air r O2 O2 = rate of oxygen consumption (mass/liquid volume time) t = time us = superficial gas velocity (length/time) V = liquid volume in the reactor
References Atkinson, Bernard, Biochemical Bernard, Biochemical Engineering and Biotechnology Handbook , Nature Press, New York, 1983, pp. 239 - 269. nd
Bailey, James E. and David S. Ollis, Biochemical Ollis, Biochemical Engineering Fundamentals, Fundamentals, 2 . ed., McGraw-Hill, New York, 1986, pp. 498-494. Dickey, David S., and Ramesh R. Hemrajani, “Recipes for Fluid Mixing”, Chemical Engineering , March 1992, pp. 82-89. McCabe, Warren L., Smith, Julian C., and Peter Harriot, Unit Operations in Chemical th Engineering , 4 . ed., McGraw-Hill, New York, 1985, pp. 208-252. Oldshue, J. Y., and N. R. Herbst, A Guide to Fluid Mixing, Mixing Mixing, Mixing Equipment Company, Company, Rochester, N.Y., 1990, pp. 1-3,. Roberts, Ronnie S., James R. Kastner, Maqsood Ahmad, and D. William Tender, “The Effect of Agitation on Oxygen Mass Transfer in a Fermentor” Chemical Engineering Education, Education, vol. 26 no. 3, 1992, pp. 142-145.
Appendix 1: Experimental Supplies Supplies and Equipment dissolved-oxygen probe Multigen fermentor Opto 22 data-acquisition system
nitrogen cylinder and regulator
Appendix 2: Details of of Experimental Procedure I.
Operation of the Multigen fermentor A. Make sure that the thermistor (a white, plastic-coated wire) wire) and resistance heating rod are properly mounted in the stainless steel tubes in the headplate.
II.
B.
Turn on the power switch, and the air, agitation, and heat switches.
C.
Adjust the temperature (°C) and impeller rate (rpm) dials to obtain the desired desired setpoint values. A thermometer can be inserted into the headplate to measure the liquid temperature (Use the ones with red liquid rather rather than mercury). The impeller rate is given by the gauge on the upper left.
Operation of the Opto 22 System A. Calibration of the Dissolved-Oxygen Dissolved-Oxygen Probe and the Opto 22 System
III.
1.
A two-point calibration will be used to convert the 4 to 20 mA signal of the oxygen amplifier to to percent-air-saturation units. Nitrogen sparging will be used to calibrate the low end (0%), and air sparging will be used to calibrate the high (100%) end. With the air and impeller rates both set high, allow the C value to reach its steady-state value.
2.
Confirm that the LED readout of the oxygen amplifier is reading 21 (the percentage oxygen in air). Instructions for calibrating the O2 sensor are located in a booklet near the Multigen operating station.
3.
Shut off the air. Connect the compressed tank to the Multigen rotameter. With the rotameter shut off, carefully open the regulator by slowly turning the bar clockwise until the pressure pressure reads about 2 psi. Then open the rotameter to initiate N2 sparging.
4.
Using high impeller and sparging rates, equilibrate the water in the Multigen with the N2 gas. The oxygen amplifier should now read a value close to zero, which will be used as the second lower calibration reading.
Experimental Runs to Determine k La A. Once the oxygen amplifier and Opto system have been calibrated, a series of runs are conducted that alternately use air or N2 as the the sparged gas. When “Start” is pressed Opto 22 save data to the end of a data file that can be opened
and manipulated by Excel as a spreadsheet. The Opto 22 display screen contains information about the location of the folder in which the file is stored. This file will contain data from previous groups” runs if it has not been erased before the current group begins to use it. The data logged during each run will later be analyzed to obtain one experimental value of k La. Because each run is fairly short, the effects of several different independent variables, such as gas flow rate and impeller rate on k La may be investigated. For the 1-L system, temperature can also be studied, although this effect is usually minor. B. Although the length of each run is only 1 to 2 min, it may be desirable to have the Opto system log data continuously for the entire lab period. C.
With the oxygen concentration within 5% of the steady-state value for the gas being sparged, set the impeller rate for the next run.
D. Turn off off the the rotameter, rotameter, and change to the other gas supply. Set the rotameter to give the desired gas flow rate, and immediately begin logging data. E.
When the CO2 value has reached about 50% saturation, enough data have been logged to accurately determine k La. Turn up the gas flow rate and impeller rate to minimize the time between runs.
F.
Once the CO2 value is within 5% of steady state, repeat the previous four steps using a different set of experimental conditions. Increments of 200 rpm and 0.5 L/min are usually sufficient for the impeller rate and gas flow rate, respectively.
Figure 1. Experimental Setup for Mass Transfer Experiment
Fermentation Power Transfer Transfer Introduction Fluid mixing is a unit operation used extensively in the chemical and biochemical processing industries. Typically, an industrial mixer consists of a motor, a gearbox, and a shaft on which one or more impellers impellers are mounted. Power is transferred from from the motor to the fluid via the shaft shaft and impeller. The power requirement, which depends on the geometry of the impeller and tank, the revolution rate, and the physical p hysical properties of the liquid, is often is an important factor in the overall economics of the process. Power requirements must therefore be either estimated using literature correlations or measured experimentally when designing a mixing process. process. The objective of this experiment is to measure the power demand of three common impeller types in a tank having standard dimensions. The experimental data are correlated using dimensionless dimensionless groups and compared to literature results.
Background There are many types of impellers, including flat-bladed turbines, marine propellers, and helical impellers. In general, impellers may may be classified as either axial-flow axial-flow or radialflow. Radial-flow impellers, such as the flat-bladed turbine, force force liquid radially outward in a direction perpendicular to the impeller shaft. Axial-flow impellers, which typically have pitched blades like marine propellers, pump the liquid in a direction parallel to the shaft. Power transferred to the liquid through the impeller is manifested as either p umping capacity (i.e., liquid flow rate through the impeller zone) or impeller head (i.e., shear shea r rate in the impeller zone). Each impeller type exhibits a characteristic balance between flow and shear. Axial-flow impellers typically typically generate high flow rates and low shear rates, whereas radial-flow impellers impellers generate more shear and less flow. Similarly, different mixing applications require different different proportions of flow flow and shear. Blending of two viscous liquids is most efficiently done with impellers having a high pumping capacity and low shear rate, while dispersion of gas in a liquid phase requires a higher shear rate and lower pumping capacity. Thus, a key objective in the design of mixing mixing systems is to match the flow properties of the impeller to the flow requirements of the application.
Experimental Equipment and Procedure The experimental apparatus consists of a cylindrical plexiglass tank fitted with removable baffles, and a Lightnin Mixer LabMaster tm tm 2510 (Lightnin Mixing Co., Rochester, NY). Three Lightnin impellers are available: available: an A100 marine propeller with with a power number of 0.87, an R100 six-bladed disk turbine with a power number of 6, and an A310 high performance, axial-flow impeller with a power number of 0.3. The LabMaster tm mixer motor measures the impeller speed and the power consumed.
Newtonian fluids having different viscosities (e.g., glycerine and water) can be tested to explore a larger range of impeller impeller Reynolds numbers. Also, the power-consumption curves of non-Newtonian fluids (e.g., aqueous polymer solutions) can be measured. Also, measurements can be made with and without gas sparging. The data should be reported in terms of dimensionless groups and compared to published results. The mixer should be auto-calibrated with the impeller shaft removed before use. Autocalibration allows the mixer to subtract the rate of po wer consumption with no impeller from the power consumption with the impeller agitating the fluid. In this way, the power data readings obtained from the mixer have been corrected for power consumption in the due to friction in the bearings and mechanical couplings.
Theoretical Analysis The correlations from Rushton, Costich, and Everett’s 1950 dimensional analysis of fluid mixing (Rushton et al., 1950) are still widely used. The key dimensionless groups for for power consumption in stirred tanks, the Power number, the impeller Reynolds number, and the Froude number, are defined below: P n = Power number =
P g c 3 i
ρ l N D
5 i
=
drag forces inertial forces
2 Di N ρ inertial forces Re = Reynolds number = = µ viscous forces 2 N i D i inertial forces Fr = Froude number = = g gravity forces
where
(1)
(2) (3)
P = power output of mixer gc = Newton’s Law unit conversion constant ρl = liquid density Ni = impeller rate Di = impeller diameter g = acceleration of gravity µ = liquid viscosity
Because the impeller tip speed is given by πDi, the numerator of the impeller Reynolds number is the product of a density, de nsity, a characteristic length, and a velocity just like other types of Reynolds numbers. The Power number can be correlated as a function of the Froude number, the Reynolds number, and ratios of the key dimensions of the tank and impeller. A general form of the correlation correlation is given below (Rushton, pt I, 1950): T t H h C c S s L l W w J j m n P n = K Re Fr ( ) ( ) ( ) ( ) ( ) ( ) ( ) Di Di Di Di Di Di Di where T = tank diameter Di = impeller diameter
(4)
H = liquid depth C = height of impeller off bottom S = pitch of impeller L = length of impeller blades W = width of impeller blades J = width of baffles B = number of blades on impeller Br = number of blades on the reference impeller R = number of baffles R r r = number of baffles on reference tank The lower-case letters are the unknown exponents of the dimensionless groups. The geometric ratios comprising the last seven terms are sometimes referred to as shape factors. Typical values of the shape factors are listed below and illustrated illustrated in Figure 2 (McCabe, Smith, and Harriot, 1985). S 1 =
C L 1 W 1 J 1 H Di 1 = = 1 S 3 = = = S 2 = S 4 = S 5 = = S 6 = = 1 T 3 T 12 T Di Di 4 Di 5
(5)
If all shape factors are constant between two mixers, the mixers are said to be geometrically similar. Under these conditions, geometric parameters parameters collapse into one constant, yielding the following simplified correlation: m
n
P n = K Re Fr
The effect of the Froude number is negligible when the tank is baffled (Rushton et al., 1950). In this case, Pn may be plotted against Re on log-log coordinates, and the slope gives the exponent of the Re term. Fluid-mixing phenomena may be classified into laminar, transition, and turbulent flow regimes in a manner analogous to pipe flow. Laminar conditions exist at impeller impeller Reynolds numbers below 10; the transition range exists between Re values of 10 and 20,000; and turbulent flow exists at Re values greater than 20,000. The Power number is analogous to the friction factor in pipe flow and the drag coefficient in flow past p ast submerged objects, in that it becomes constant c onstant in the turbulent regime for certain systems (e.g., baffled tanks with radial-flow radial-flow impellers containing Newtonian fluids). This constant value is sometimes reported by manufacturers as the p ower number of the impeller.
(6)
Symbols
µ = liquid viscosity ρl = liquid density B = number of blades on impeller Br = number of blades on the reference impeller C = height of impeller off bottom Di = impeller diameter Di = impeller diameter 2 Fr = Froude number = Ni Di/g g = acceleration of gravity gc = Newton’s Law unit conversion constant H = liquid depth J = width of baffles L = length of impeller blades Ni = impeller rate P = power output of mixer 3 5 Pn = Power number = Pgc/(ρ N l i Di ) R = number of baffles Re = Reynolds number = D N i ρ/µ R r r = number of baffles on reference tank S = pitch of impeller T = tank diameter W = width of impeller blades
References Bailey, James E. and David S. Ollis, Biochemical Ollis, Biochemical Engineering Fundamentals, Fundamentals, 2 nd. ed., McGraw-Hill, New York, 1986, pp. 498-494. Dickey, David S., and Ramesh R. Hemrajani, “Recipes for Fluid Mixing”, Chemical Engineering , March 1992, pp. 82-89. McCabe, Warren L., Smith, Julian C., and Peter Harriot, Unit Operations in Chemical th Engineering , 4 . ed., McGraw-Hill, New York, 1985, pp. 208-252. Oldshue, J. Y., and N. R. Herbst, A Herbst, A Guide to Fluid Mixing , Mixing Equipment Company, Rochester, N.Y., 1990, pp. 1-3,. Rushton, J. H., Costich, E. W., and a nd H. J. Everett, “Power Characteristics of Mixing Impellers, Part I” Chemical Engineering Progress, Progress, vol. 46 no. 8, August 1950, pp. 395404. Rushton, J. H., Caustic, E. W., and an d H. J. Everett, “Power Characteristics of Mixing Impellers, Part II” Chemical Engineering Progress, Progress, vol. 46 no. 9, Sept. 1950, pp. 467476.
Appendix 1: Experimental Supplies Supplies and Equipment Equipment Lightnin Labmaster TS2510 three impellers plexiglass tank with removable baffles Camile Data Acquisition and Control system Thermometer Water Glycerine Water-soluble polymer
Appendix 2: Details of of Experimental Procedure I.
Determination and of Tank Shape Factors A. Measure all pertinent tank, baffle, and impeller impeller dimensions. B.
Identify each of the three types of impellers. The A310, A100, and R100 impellers have diameters of 3.4 in, 2.7 in, and 4 in, respectively. Verify these measurements.
C.
Based on typical values of the shape factors, choose the height above the tank bottom the impellers should be located and the total desired liquid depth in the tank.
II.
Autozeroing the Labmaster Mixer A. The mixer mixer must be raised above the tank to remove the shaft. Locate the two hand-tightened knobs and the metal positioning pin that hold the mixer in position on its vertical track. With the mixer off, loosen the two knobs, and slide the mixer up the track track about 18 inches. Move the positioning pin up to the next hole. Rest the mixer on the the positioning pin, and then hand-tighten the the two knobs. B.
Depress the rectangular shaft-lock button located at the bottom center of the mixer while gently rotating the the shaft chuck with your fingers. Within half a rotation, the button should engage and lock the shaft shaft in place. While depressing the button, loosen the chuck by turning it counterclockwise (as viewed from below) until the shaft glides freely out of the mixer.
C.
Note: Never attempt to stop the mixer with the shaft lock button. Instead, use the RUN/STOP button to stop the mixer . The RUN/STOP button will have no effect if the mixer is being controlled by the computer. In this case, the power button may be used as an emergency shut-off.
D. Simultaneously press the ON and RESET buttons. Then release the RESET button before the ON/OFF button. The LCD readout should say “diagnostic” at
this point. Press “6” and then ENTER. The mixer will autocalibrate itself at different speeds and then stop. This process takes takes about 1 min. E.
Mount the desired impeller impeller on the bottom of the shaft, making sure the set screw screw is snugly tightened. Mount the R100 impeller with the hub facing up. Mount the A100 and the A310 impellers with the convex side of the blades facing up. When properly installed, the A100 and A310 should pump liquid downward.
F.
With the mixer stopped, slide the shaft into the mixer so that the end of the shaft extends to the top of the mixer. mixer. Depress the shaft-lock button, and snugly handtighten the chuck by turning it clockwise (as viewed from below). Please do not use tools to tighten the chuck.
G. Move the positioning pin to its lowest position. Then loosen the two knobs, and lower the mixer until it rests on the positioning positioning pin. Tighten the two knobs. H. Press the ON/OFF ON/OFF button twice to get the normal operating operating mode. III. Measurement of Power Consumption A. Use the MENU key to move the cursor (i.e., flashing asterisks) asterisks) from speed to the temperature/power window. Press the ITEM key several times in succession to show the air temperature temperature and the power in Watts. Watts. Press the MENU key to move the cursor to the pumping capacity setting, and use the ITEM key to select the impeller impeller type used. When prompted, enter the impeller diameter to one decimal place. B.
Locate the impeller impeller at the proper depth in the tank by sliding the shaft up and down in the chuck. However, at all times the top of the shaft should be at or above the top of the mixer. If necessary, the impeller may may be loosened and slid down the shaft. It may be helpful to raise raise the mixer to adjust the shaft. The knobs that hold the mixer in the track must be tightened before operation.
C.
Fill the tank to the the desired desired depth with the test test liquid. liquid.
D. Measure the power consumption rate over a wide range of rpm values: about 50 to 400 rpm for the R100 and 50 to 800 rpm for the the other two impellers. Allow the system to equilibrate at each speed. Also, measure the temperature of the liquid with a thermometer (The temperature value given by the mixer is the air temperature). It is important to watch the impeller and shaft carefully at all times during operation for any signs of slippage. If the impeller or shaft appear to be going up or down while rotating, immediately stop the mixer. Also, after stopping the mixer, check for evidence of scoring due to slippage of the shaft. If slippage is suspected, consult the instructor. E.
Raise the mixer, mixer, and empty empty the the tank. tank. Lower the impeller impeller into the empty empty tank, and measure the power consumption rate of the impeller in air for the same rpm
values. Most of these values will be zero, so start at the upper end of the rpm range.
Figure 1. Tank and Impeller Constants
Membrane Filtration Introduction Membrane separations are frequently used in the growing biotechnology industry. Polymer membranes separate large molecules or particulates from a liquid stream based on molecular size. Substances smaller than the pore size of the membrane pass through the membrane with the solvent; solvent; this fraction is called the permeate. The fraction that does not pass through the membrane is called the retentate. Filtration processes may be categorized by pore size. Microfiltration membranes membranes have pores ranging from 0.01 to 10 microns in diameter and are commonly used to separate whole cells from fermentation fermentation broth. Ultrafiltration membranes have pores 10 to 100 A in diameter, which correspond to molecular weight cutoffs of 500 to 1,000,000 daltons. These membranes are often used to remove macromolecules, such as proteins, peptides, and viruses. Reverse osmosis (RO) membranes have even smaller pores that that can retain salts while passing passing water. They are generally used for water purification purposes (Tutunjian, 1985). The objective of this experiment is to analyze a nalyze the performance properties of two types of filters commonly used in in biotechnology applications: a hollow-fiber microfilter microfilter and a spiral-wound ultrafilter. The microfilter will be used to concentrate a suspension suspension of yeast cells, and the ultrafilter ultrafilter will be used to concentrate a blue dextran solution. The membrane flux and rejection coefficient will be measured as a function of transmembrane pressure difference. The intrinsic and concentration-polarization resistances of the membrane will also be determined.
Background Two types of filtration filtration exist: dead-end and crossflow. crossflow. Dead-end filtration produces a cake that builds up on the membrane, reducing reducing the filtration rate as it grows. Crossflow filtration avoids cake formation by passing the fluid rapidly across the surface of the membrane. The resulting fluid shear sweeps sweeps away particles that might might otherwise build up on the membrane surface (Murkes, 1988). 1988). Both types of filtration filtration are illustrated schematically in Figure 1. Commercially available membrane geometries may be classified into two categories: hollow fibers and flat sheets. Hollow fibers are self-supporting, self-supporting, porous tubes contained in cartridges. Hollow fibers are able to to be backflushed, facilitating cleaning and product recovery. Flat-sheet membranes exist in many many forms, including conventional plate-andframe filters and spiral cartridges. In general, flat sheet membranes membranes can operate at higher pressures but can be difficult to clean (Tutunjian, 1985). Membrane filtration can be used to concentrate, diafilter, or purify a solution or suspension. Concentration is achieved by removing a portion portion of the solvent through the membrane while retaining larger components. In diafiltration, low-molecular-weight solutes (e.g., salts) are removed from solution by exchang ing them for water across the
membrane, while the higher-molecular-weight product is retained. Purification is the isolation of the final product from other solutes (Tutunjian, 1985). The advantages of membrane filtration over conventional separation methods are numerous. Membrane processes operate at low temperatures, thus allowing separation separation of heat-sensitive products and minimizing energy costs costs (Murkes, 1988). Membrane separations also facilitate other downstream-processing steps. steps. For instance, because the filtrate is typically free of colloids and microorganisms fouling of chromatography resins is reduced. If the products are retained retained by the membrane, they may be purified and concentrated in one step. Products permeable to the membrane are separated separated from larger molecular weight substances with minimal dilution. Despite its utility, utility, membrane filtration also poses some disadvantages. Low membrane filtration rates are common, requiring relatively large membrane areas and therefore a substantial capital investment. Also, membranes membranes may foul. Many common anti-foaming agents used in fermentations adhere to the membrane and rapidly plug it (Murkes, 1988). The shear inherent in crossflow filtration may damage shear-sensitive products (e.g., proteins and mammalian cells). High-viscosity solutions, such as mycelial fungi fermentation broths require high pumping rates to minimize fouling a nd maintain a suitable flux (Cheryan, 1986). Concentration polarization, the formation of a ge latinous layer on the surface of the membrane, increases the resistance resistance to flow and lowers the membrane membrane flux. This effect can be minimized by operating the membrane at high crossflow velocities to minimize the thickness of the layer (McGregor, (McGregor, 1986). In some cases, concentration polarization polarization has reduced the flux by 95% within 25 seconds (Cheryan, 1986).
Experimental Equipment and Procedure The spiral-wound ultrafiltration system and hollow-fiber microfiltration system used in this experiment are shown shown schematically in Figures 2 and 3. Each system includes a liquid reservoir, a membrane module, a peristaltic (tubing) pump, and valves to control the pressure difference difference across the membrane. The Amicon H1P100 hollow-fiber hollow-fiber cartridge (Danvers, MA) has a molecular weight cutoff of 100,000 daltons. It contains 55 2 fibers, each having a 1.1 mm inner diameter, giving a total membrane area of 0.3 m . The Amicon S1Y30 spiral cartridge has a molecular-weight cutoff of 30,000 daltons, and 2 a membrane area of 0.0929 m . The hollow-fiber cartridge cartridge is used to concentrate a suspension of yeast, and the spiral-wound ultrafilter is used to concentrate blue dextran. The experiment is initiated by loading the reservoir with a known volume of the solution to be concentrated. The liquid is then pumped through the the membrane cartridge at a measured rate. The back-pressure valve is partially partially closed to restrict the flow downstream of the membrane, and thus generate a pressure difference between the filtrate and permeate sides of the membrane. membrane. The volumes and solute concentrations of both the permeate and retentate are then recorded as a function of time. These data are later analyzed to determine the effects of transmembrane pressure drop and liquid flow rate on the rejection coefficient and membrane flux.
A spectrophotometer or colorimeter is used to determine the concentrations of the solutes (yeast cells or blue dextran) by their optical densities densities (OD) at 600 nm. Optical density, also referred to as absorbance, is directly proportional to the concentration of a substance that either absorbs or diffracts light for for OD values below about 1.0. In this experiment, OD measurements will be directly used as a c oncentration unit, without converting them to more conventional concentration units, such as g/L.
Theoretical Analysis Transport Models The net pressure driving force (P) driving liquid transport through the membrane pores is the difference between the transmembrane hydrostatic pressure difference (∆PT) and the osmotic-pressure difference (∆π): P = ∆ P T - ∆π
(1)
For dilute solutions and separation of macromolecules, such as proteins, the osmotic pressure difference is usually small compared to the hydrostatic pressure pressure difference. Under these conditions, P may be taken as equal to ∆PT. However, for many reversereverseosmosis processes (e.g., water purification from brine), the osmotic pressure difference is quite large. The rate of liquid transport across the membrane is usually expressed as a flux (J), which has units of mass (or moles) per unit cross-sectional area of membrane membrane per unit time. The flux varies with the net driving force as well as the physical properties of the membrane, the solvent and the solute. At relatively small values of P, J is directly directly proportional to P. However, for large values of P, J is essentially independent of P. Mathematical models have been developed for both of these regimes, as well as the intermediate regime. At low operating pressures, the membrane flux may be described by the Hagen-Poiseuille law for laminar flow through channels, as shown b elow: J =
ε r 2 P 8 µ L p
where J = membrane flux r = channel (mean pore) radius P = net pressure driving force µ = fluid viscosity L p = membrane thickness (or pore length) ε = surface porosity of the membrane This model is based on assumptions of steady state, laminar flow through the pores, an incompressible, Newtonian fluid, and negligible end effects. It predicts that flux should be directly proportional to the transmembrane pressure difference and inversely proportional to the viscosity. The viscosity at the membrane surface may differ from that in the bulk fluid due to the buildup of excluded solute molecules at the membrane surface. This buildup is
(2)
frequently referred to as concentration polarization. Thus, flux is directly proportional to pressure only when concentration polarization is minimized. This condition exists when the feed composition is low, the transmembrane pressure difference is low, and when the crossflow velocity is high (Cheryan, 1986). At high operating pressures, a gelatinous layer, referred to as the concentration polarization layer, develops at the membrane surface. As the layer thickens, the flux becomes relatively insensitive to the transmembrane pressure difference. difference. Film theory can be effectively used to describe membrane performance in this regime. Figure 4 shows the two films that are assumed to provide mass-transfer resistances in series at the membrane surface: the concentration-polarization layer and the liquid boundary layer. The continuous flow of permeate through the membrane causes solute molecules to be carried from the bulk solution to the membrane at a rate given by the following equation: J s = J X b
(3)
where Js = convective mass flux of solute toward the membrane J = permeate flux X b = bulk concentration of the retained (rejected) solute When flow is first initiated, this convective flux results in the accumulation of solute at the membrane surface, leading to a higher solute concentration at the membrane surface than in bulk solution. The resulting concentration gradient induces induces a diffusive flux of solute in the opposite direction. direction. At steady state, the convective flux is exactly counterbalanced by diffusive flux. This relationship may be expressed as shown below, using Fick’s law to describe the diffusive flux: J s s = - D
dC dx
(4)
where D = diffusion coefficient dC/dx = concentration gradient in the boundary layer Eliminating Js from Equations 3 and 4, and integrating across the boundary layer of thickness δ gives the following result (Cheryan, 1986): J =
D
C g C g ln( ) = k ln( ) δ C b C b
where k = mass transfer coefficient δ = film (boundary layer) thickness Cg = solute concentration at the membrane surface C b = concentration solute in the bulk fluid
(5)
This model predicts that the flux is controlled by the rate of solute transfer back into the bulk phase, and that the flux is independent of pressure. Thus, the high-pressure regime, where the permeate flux is unaffected by pressure, is also known as the “mass-transferlimited” regime. The concentrations Cg and C b are fixed, so to increase the flux in this regime, the mass-transfer mass-transfer coefficient must be increased. This may be done by increasing the crossflow velocity, which increases the shear rate a t the membrane surface, and thus reduces the boundary layer layer thickness. Mass-transfer coefficient data have been correlated correlated using dimensionless groups, as shown below (Cheryan, 1 986): α β d ω Sh = A (Re ) (Sc ) ( h ) L
(6)
where dh = hydraulic diameter L = channel length Sh = Sherwood number = kdh/D Re = Reynolds number = ρudh/µ Sc = Schmidt number = µ/ρD D = diffusion coefficient µ = fluid viscosity ρ = fluid density u = fluid velocity α,β,ω,A = empirical constants The hydraulic diameter is defined as 4 times the cross sectional area divided by the wetted perimeter. For tubes and circular circular cross sections, sections, dh = diameter (d). (d). An example correlation for fully developed laminar flow is 0.33
Sh = 1.86 Re0.33 Sc0.33 ( d h /L )
(7)
An example correlation for turbulent flow with Re > 4000 is (Cheryan, 1986): Sh = 0.023 Re0.8 Sc0.33
(8)
Correlations such as these allow k, and hence J, to be predicted for a membrane operated in this regime. A third model used to predict membrane fluxes combines the features of the two previous models. This model is based on the common theme in transport phenomena that flux is directly proportional to the driving force and inversely proportional to the sum of the resistances. The Hagen-Poiseuille equation may may be expressed in such a form: J =
P R m′
where R m = intrinsic membrane resistance
(9)
Fouling of the membrane occurs when whe n some of the pores get partially pa rtially or fully plugged with particulates. The resistance due to fouling is generally unaffected by the pressure pressure drop and is thus indistinguishable from R m. For convenience, then, then, these two resistances resistances ’ may be lumped together and renamed R m . Because this lumped lumped resistance and the resistance due to concentration polarization are a ssumed to act in series, they may be summed, as shown below: J =
P
(10)
Rm′ + R p
where R m’ = intrinsic membrane resistance including fouling R p = resistance due to concentration polarization As discussed previously, the resistance due to concentration polarization increases with P. As a first estimate, estimate, the resistance may may be assumed to be proportional to P: R p = φ ∆ P T
(11)
where φ is an empirical constant that reflects the degree of concentration polarization. Combination of the Equations (47), (48) and (49) results in the following equation (Cheryan, 1986): J =
P Rm′ + φ P
This model can predict the experimental trends observed in both the high-pressure and low-pressure regimes. At low P, the first first term in the denominator dominates, and the flux is proportional to P. The filtration rate may may be controlled in this regime regime by adjusting operating pressure. At high P, the the second term dominates, and the flux flux is a constant. The filtration rate in this regime may be controlled by adjusting the crossflow velocity, which decreases the concentration polarization polarization and boundary layer thicknesses. Crossflow velocity has an especially strong effect on the flux of systems with particulates, such as bacteria (Tutunjian, 1985). At intermediate P values, transitional behavior is observed. The experimental data may be analyzed by determining values of R m’ and φ that allow Equation 12 to reproduce the experimental data. Equation 12 has the same form as the Michaelis-Menten equation. Consequently, the double-reciprocal plot approach (1/J vs 1/P) would enable R m’ m’ and φ to be determined graphically. The graphical method is described in more detail in the Enzyme En zyme Kinetics chapter of this manual .Inspection of the data should also indicate whether the membrane systems are operating in the low pressure, intermediate, or high-pressure regime. Material-Balance Models The material-balance models are not concerned with transport rates. Instead, they establish relationships between the volumes and concentrations of the various streams in
(12)
the filtration process. A key parameter used to describe describe the membrane’s ability to retain the solute is the rejection coefficient (σ). The rejection coefficient is the probability probability of a solute molecule being excluded when it encounters a pore. Thus, σ is related to the probability (p) of the molecule passing through the pore by the following relation: σ = 1 - p
(13)
The value of σ may be experimentally determined from instantaneous measurements of the solute concentration in the reservoir (feed) stream and the permeate stream using the following identity: σ = 1 -
C p C
(14)
where C p = solute concentration in the permeate C = solute concentration in the reservoir reservoir A rejection coefficient of 1 (i.e., p = 0) indicates that the solute is completely retained, while a rejection coefficient of 0 (i.e., p = 1) implies that the solute freely passes through the membrane with the solvent. solvent. The value of p is assumed to to be constant for most filtration processes. Different material-balance models exist for different types of membrane processes. The nomenclature for the models that follow is illustrated illustrated conceptually in Figure 5. In all cases, the recycled retentate stream is assumed to mix instantaneously with the liquid in the reservoir. In diafiltration, unwanted low-molecular-weight solutes are removed by exchanging them for water molecules across the membrane. Typically, the solutes removed are salts salts that were used as buffers or to precipitate proteins out of aqueous solution (Tutunjian, 1985). The concentration of the removed solute decreases exponentially with the volume of water exchanged, according to the following equation: C = C o exp(- V d / V o ) where C = solute concentration in the reservoir reservoir Co = initial solute concentration Vd = total volume of water exchanged Vo = initial volume of solution in the reservoir However, the concentration of the higher molecular-weight solutes remains essentially unchanged in diafiltration. Purification processes are similar in operation to diafiltration; however, in purification the smaller, removed solute is the desired product. The recovery (R), defined as the the amount
(15)
of the solute in the permeate divided by the amount of o f solute initially present, may be calculated using the following equation:
R =
V p C p V o C o
=1-
V C
(16)
V o C o
where Co = initial solute concentration in the reservoir V = volume of retentate V p = permeate volume In a batch concentration process, a volume of permeate (V p) is removed through the pores of the membrane, decreasing the retentate volume from the initial value (Vo) to its final volume (V). If the liquid held in the membrane pores is neglected, these volumes volumes are related by the simple material balance = V o - V p
(17)
The volume-concentration ratio (VCR) model is frequently used to describe the performance of membrane-based concentration processes. The VCR is simply the ratio of the initial liquid volume to the final liquid volume: VCR =
V o V
(18)
As described by Cheryan (1986), the VCR model can be developed from material balances on an incremental volume of liquid (∆V) passing through the membrane. The incremental amount of solute (∆ N) passing through the membrane is given by
∆ N = (C ∆V) p
(19)
Dividing both sides of this equation by the total amount of solute in the retentate (N), substituting in the identity C = N/V, and then taking the limit as ∆V goes to zero yields
dN N
= p
dV V
(20)
Integrating and rearranging gives
N = KV P
(21)
where K is an integration integration constant. Dividing by V gives a similar similar expression in terms terms of the reservoir concentration: C = K V p-1
(22)
Rewriting p in terms of σ, and eliminating K with the following identity
K =
C -σ
V
=
C o
(23)
-σ
V o
gives the following equation: σ
C = C o (VCR )
(24)
This equation predicts that a logarithmic plot of C versus VCR should yield a straight line with a slope of σ. Deviation from this trend trend would suggest that the solute solute was being adsorbed by the membrane or that the rejection coefficient of the membrane was varying with concentration. The yield (Y) of the concentration conce ntration process may be defined as the mass of solute present in the retentate divided by the mass of the solute initially present:
Y =
C V C o V o
σ -1
= (VCR )
This yield is analogous to the recovery (R) of the diafiltration process.
Symbols
δ = film (boundary layer) thickness ∆π = osmotic pressure gradient α,β,ω,A = constants ∆PT = transmembrane pressure drop ε = surface porosity of the membrane µ = fluid viscosity ρ = fluid density φ = empirical constant reflecting the degree of concentration polarization B = membrane permeability coefficient C = concentration of the solute in the retentate C b = bulk concentration of the retained (rejected) solute Cg = solute concentration at the membrane/gel interface Cm = retentate concentration at the membrane surface Co = initial solute concentration in the reservoir C p = solute concentration in the permeate CS = concentration of removed solute
(25)
CSo = initial concentration of removed solute D = diffusion coefficient dC/dx = concentration gradient over a differential element in the boundary layer dh = hydraulic diameter = 4(cross sectional area)/(wetted perimeter) J = permeate flux, mass (or moles)/area⋅time Js = rate of solute transport by convection, mass (or moles)/area⋅time k = mass transfer coefficient L = channel length Lc = length required for a fully developed concentration profile L p = membrane thickness (or pore length) Lv = length required for a fully developed velocity profile N = mass (or moles) of solute in the reservoir P = net driving force for an ideal membrane r = channel (mean pore) radius R = fractional solute recovery in a purification process Re = Reynolds number = ρudh/µ R m = intrinsic membrane resistance for unfouled membrane R m’ = resistance of membrane including fouling R p = resistance due to concentration polarization = φP Sc = Schmidt number = µ/ρD Sh = Sherwood number = kdh/D u = fluid velocity V = volume of the liquid in reservoir Vd = total volume of water added Vo = initial liquid volume in reservoir V p = permeate volume X b = mass fraction of solute Y = fractional solute yield of a concentration process
References Bailey, J.E. and Ollis, D.F.(1986) Biochemical D.F.(1986) Biochemical Engineering Fundamentals, Fundamentals, pp. 100-130, McGraw-Hill Book Co., New York pp. 726 72 6 -. Beaton, N. C., “The Application of Ultrafiltration to Fermentation Products”, Polymer Products”, Polymer Science and Technology, Volume 13: Ultrafiltration Membranes and Applications, Applications, Plenum Press, New York, 1980, pp. 381 -385. Cheryan, Munir, Ultrafiltration Handbook , Technomic Publishing Co., Lancaster, Pennsylvania, 1986, pp. 74-120, 280-284. Meireles, Martine, Pierre Aimar, and Victor Sanchez, “ Albumin Denaturation During Ultrafiltration: Effects of Operating Conditions and Consequences on Membrane Fouling,” Biotechnology and Bioengineering , vol. 38, pp. 528-534, 1991. McGregor, W. Courtney, ed. , Membrane , Membrane Separations in Biotechnology, Biotechnology, Marcel Dekker, Inc., New York, 1986, pp. 61-79.
Murkes, Jakob, and Claes-Goran Carlsson, Crossflow Filtration: Theory and Practice, Practice, John Wiley and Sons, New York, 1988, pp.1-3, Tutunjian, Robert S., “Scale-up Considerations Con siderations for Membrane Processes”, Biotechnology, Biotechnology, vol 3, July 1985, pp. 615 - 626.
Appendix 1: Equipment and Reagents Needed Needed Yeast cells in solution Cell growth medium glucose yeast extract NaOH solution hollow-fiber membrane apparatus spiral-wound membrane apparatus Blue Dextran (M.W. 200,000) spectrophotometer
Appendix 2: Details of of Experimental Procedure: I.
Preparation of Yeast Culture A. About 500 mL of yeast growth medium should be autoclaved, cooled, and inoculated during the first laboratory period, so the cells will be ready by the second period. The medium contains 2% glucose (dextrose), 2% peptone, and 1% yeast extract. Titrate the medium to a pH of 7.0 with NaOH. Then, autoclave the medium in a 1 L Erlenmeyer flask vented at the top with a foam plug. Detailed instructions for operating the autoclave are given in another section of this manual. B. When the medium has cooled to about 30°C, aseptically withdraw and refrigerate a 10 mL sample to to serve as a spectrophotometer blank. Then, inoculate the medium with yeast working culture from the refrigerator. refrigerator. Add about 2 mL of working culture for every 100 mL of medium (i.e., use a 2% inoculum). C. Incubate the inoculated medium at 30 to 35°C. Arrange for the cells to be removed from the incubator in about 24 h and refrigerated.
II.
Determination of Solute-Free Flux A. Fill the reservoir with fresh RO water. Completely open the the backpressure backpressure valve and the valve at the bottom of the liquid reservoir (the handle should be in line with the pipe). Note: failure to open the backpressure valve before turning on the pump could result in high pressures that could rupture the membrane. B. Turn the pump switch to the FORWARD position. The liquid liquid should should flow flow from the reservoir, through the pump, through the membrane module, and then back into the reservoir through the the liquid-return tube. If it flows in the opposite
C.
D.
E.
F.
direction, turn the pump switch to to REVERSE. With the backpressure valve fully open, flush the membrane to rinse out any residues. If significant residues come out of the membrane, repeat the process with fresh RO water. Set the pump at the first first flow flow rate to be tested. If the pressure reading of the lower (inlet) pressure gauge is more than 4 psi higher than the upper (outlet) gauge, reduce the flow rate. Gradually close the backpressure control valve to the increase the transmembrane pressure drop to the desired value (up to about 8 psi for the hollow-fiber membrane and 30 psi for the spiral membrane). Measure the volumetric volumetric flow flow rate of the the permeate and retentate streams by directing the flow into a graduated cylinder for a measured period of time. The membrane flux can be calculated by dividing the mass flow rate of the permeate by the cross-sectional area of the membrane. Repeat the process for other flow rates and pressure drops. Each run takes about 15 min.
III. Concentration of Blue Dextran using using Spiral-Wound Ultrafiltration Membrane A. Empty the liquid liquid from the reservoir, tubing, and membrane membrane module. If the liquid contains sodium azide, save it in a labelled container, and reuse it when the day’s experiments are done. Warning: sodium azide is a potent poison. Wear gloves when handling it, and clearly label any reservoir that contains it. B. Pour 2 L of RO water into the reservoir, and, with the backpressure valve fully open, pump the water through the membrane module to flush out the system. Gradually increase the backpressure control valve to flush some of the RO water through the permeate tube. Keep the backpressure below 30 psi at all times. Remove the RO wash water from the tank, membrane module, and tubes, and pour it down the drain. C. Pour 1 to 2 L of relatively dilute (about 0.1 g/L) g/L) blue dextran solution into the reservoir. The blue dextran solution of a previous group should be used, if possible, because it is quite expensive. Blue dextran solutions should be stored in the refrigerator when not being used to prevent microbial growth. D. Set the flow rate to the desired value. If the pressure reading of the lower (inlet) pressure gauge is more than 4 psi higher than the upper (outlet) gauge, reduce the flow rate. E. Gradually close the backpressure valve until the desired transmembrane pressure difference is obtained (up to about 30 psi). When all air has been displaced from the permeate tube, the experiment is ready to begin. F. To initiate initiate the experiment, place the end of the permeate tube into a graduated cylinder and begin timing. At regular time intervals, intervals, record the time, pressures, and volumes of the liquid in the reservoir and graduated cylinder. Also, measure the OD (600 nm) of the samples taken from the retentate stream, the permeate stream, and the reservoir at regular intervals. The mathematical model is based on the assumption that the reservoir contents are well mixed. Therefore, it would be good practice to stir the reservoir occasionally, especially just before sampling the reservoir. The retentate and permeate samples should be taken at the tube outlets.
G. When the run is complete, recombine the permeate and filtrate, filtrate, and repeat the experiment at a different flow flow rate or pressure drop. Each run takes about 10 min. H. When the day’s runs are finished, fully open the backpressure valve, and drain the reservoir, tubes, and membrane module. I. Refill the tank with reverse osmosis (RO) water, and flush flush out the entire entire system, including the permeate tube. J. Drain the system, and repeat the flushing procedure again. K. Replace the RO water with an antimicrobial agent consisting of a 0.2% aqueous solution of sodium azide. Pump this solution through the system, including the permeate line. Clamp off the end of the permeate tube, and submerge the end of the liquid-return tube in the liquid in the reservoir to prevent evaporation. Warning: sodium azide is a potent poison. Wear gloves when handling it, and clearly label any reservoir that contains it. Reuse the sodium azide solution whenever possible. IV. Concentration of Yeast Cells using Microfiltration Microfiltration Membrane A. Warm the yeast culture up to room temperature. B. Empty the liquid from the reservoir, tubing, and membrane membrane module. If the liquid is sodium azide solution, save it in a labelled container, and return it to the membrane system when the day’s experiments are done. Warning: sodium azide is a potent poison. Wear gloves when handling it, and clearly label any reservoir that contains it. C. Pour 2 L of RO water into the reservoir, and, with the backpressure valve fully open, pump the water through the membrane module. Gradually increase the backpressure control valve to flush some of the RO water through the permeate tube. Keep the backpressure below 8 psi at all times. Remove the RO wash water from the tank, membrane module, and tubes, and pour it down the drain. D. Pour the yeast culture into the reservoir, and dilute to about 2 L with RO water. E. Gradually close the backpressure valve until the desired transmembrane pressure difference is obtained (up to about 8 psi). Note that the hollow fibers cannot withstand as large a transmembrane pressure difference as the spiral membrane. When all air has been displaced from the permeate tube, the experiment is ready to begin. F. To initiate initiate the experiment, place the end of the permeate tube into a graduated cylinder and begin timing. At regular time intervals, intervals, record the time, pressures, and volumes of the liquid in the reservoir and graduated cylinder. Also, measure the OD (600 nm) of the samples taken from the retentate stream, the permeate stream, and the reservoir at regular intervals. The mathematical model is based on the assumption that the reservoir contents are well mixed. Therefore, it would be good practice to stir the reservoir occasionally, especially just before sampling the reservoir. The retentate and permeate samples should be taken at the tube outlets. G. When the run is complete, recombine the permeate and filtrate, filtrate, and repeat the experiment at a different flow rate or pressure drop. H. When the day’s runs are finished, fully open the backpressure valve, and drain the reservoir, tubes, and membrane module.
I.
Refill the tank with reverse osmosis (RO) water, and flush flush out the entire entire system, including the permeate tube. J. Drain the system, and repeat the flushing procedure again. K. Replace the RO water with an antimicrobial agent consisting of a 0.2% aqueous solution of sodium azide. Pump this solution through the system, including the permeate line. Clamp off the end of the permeate tube, and submerge the end of the liquid-return tube in the liquid in the reservoir to prevent evaporation. Warning: sodium azide is a potent poison. Wear gloves when handling it, and clearly label any reservoir that contains it. Reuse the sodium azide solution whenever possible.
Figure 1. Schematic Comparison of Dead End and Crossflow Crossflow Filtration
Figure 2. Diagram of Spiral Cartridge Apparatus
Figure 3. Diagram of Hollow Hollow Fiber Apparatus
Figure 4. Schematic Concentration Concentration Polarization Polarization and Boundary Layer
Figure 5. Schematic Diagram of Membrane Membrane System Showing Nomenclature Nomenclature
Operating Procedures for Autoclave and Spectrophotometers I.
Autoclave A. Please coordinate the autoclaving runs with with other groups to minimize the runs needed and maximize everyone’s efficiency. Each run takes about an hour. B. The chamber of the autoclave is easily recognized by its strong door with a large handwheel used to crank tighten the chamber door. To the left of the chamber is a small door below a silver “warning” plate. If the door is not already down, slide it down, and turn on the POWER and CONTROL switches. The control panel will then light up. C. Set the the STERILIZE time thumbwheel to 15 minutes and the dry time time thumbwheel to 0 min. The cycle time will be displayed displayed on the primary control panel. Once the cycle has been started, the time cannot be changed using the thumbwheels. D. Make sure that the thumbwheel above the printer is set set to 121 for 121°C. If the temperature is outside the allowable range, a buzzer will sound until an allowable temperature is selected. Once the cycle has started, the temperature cannot be changed using the thumbwheels. E. Open the upper access door located just just above the chamber. Turn the steam steam and water valves all the way on (counterclockwise). (counterclockwise). Make sure that the chamber pressure gauge reads near 0 psi (gauge) and the manual control knob is turned to “off”. The HI-LO knob should already already be set to LO and should not need adjusting. F. If the pressure in in the chamber is sufficiently low (about 0.5 psig), the chamber door may be opened by turning the handwheel counterclockwise. If the door will not unlock, try turning the handwheel ha ndwheel slightly clockwise and press the center black button on the the wheel. The autoclave has a safety feature that prevents the handwheel from turning if there is significant steam pressure in the chamber. G. Make sure that all containers to be sterilized sterilized are properly vented to allow allow steam to freely enter and exit. Vessels should have at least one cotton, foam, or a membrane sterile barrier to allow gas flow while while maintaining sterility. Place the items to be sterilized in one of the plastic autoclaving trays, and load the tray into the chamber. Close the door, and press on the door to ensure a good seal. H. Turn the handwheel clockwise to lock the door. The DOOR DOOR UNLOCKED UNLOCKED light on the panel should shut off at this time. Once the chamber is pressurized, an integral pressure lock will prevent the door from being opened. I. Press the liquids cycle selector on the lighted panel. It It should increase in brightness. The CONDITION and WARNING HOT LIQUIDS lights should come on. If you press the wrong cycle selector selector button, simply press the white reset button to stop the cycle, and then depress the liquids button.
J.
K.
L.
M.
N.
II.
After the CONDITION phase is completed, the STERILIZE light comes on for the duration of the sterilization sterilization phase. The STERILIZE TIME readout begins to count down once the target temperature and pressure pressure have been reached. The printed record will show when the sterilize time began. When the sterilization is complete, complete, the EXHAUST light comes on, and the chamber slowly begins to depressurize. The printed record will show show the time, temperature and pressure when the exhaust phase begins. Upon completion completion of the exhaust phase, the buzzer will sound, and the COMPLETE light will come on. Crack open the door about ½”, and leave it cracked for about 10 minutes. Do not fully open the door at this time. When the door is opened the the buzzer will stop and the WARNING HOT LIQUIDS light will begin to flash. At the end of ten minutes, minutes, the buzzer will sound again, and the WARNING HOT LIQUIDS will stop flashing but remain lit. Press the reset reset button to silence the buzzer. Open the chamber door, and carefully remove the load from the sterilizer using the insulated gloves. Press the reset button to shut off the warning warning hot liquids light, and tear off the paper form form the printer. Shut off the power and control switches for the sterilizer, and shut off the steam and water.
Perkin-Elmer Lambda 3A Spectrophotometer A. The spectrophotometer is a delicate and expensive electronic instrument. Do not use containers with liquids on or above the spectrophotometer. B. Turn on the spectrophotometer, spectrophotometer, and allow it to warm warm up for 15 min. Make sure the appropriate lamp is on for the wavelength range of interest. Below 300, use the UV lamp; between 300 and 360, use both; and above 360, use the visible lamp. C. Type in the desired wavelength (in nm) on the key pad, and press the GO TO key. D. Fill two clean cuvettes with the blank solution (e.g., distilled distilled water). water). Place one in the rear (blank) cuvette holder and the other in the front (sample) holder. If UV wavelengths (less than 400) are to be used, cuvettes designated as “UV grade” should be used. Otherwise the cuvette windows may absorb too much UV light, reducing the sensitivity of the assay. E. Zero the instrument by pressing the ZERO key. F. Empty the sample sample cuvette, taking care not to get any liquid on the outside outside of the clear glass windows. Rinse it with distilled distilled water, and gently touch the mouth of the inverted cuvette to a paper towel to siphon off any remaining water. You may wish to find cuvettes with identical ide ntical absorbances. G. Transfer the sample into the sample cuvette. Insert it into the spectrophotometer, and close the cover. Record the OD. H. To analyze another sample, repeat the previous two steps.
IV. Jenway Colorimeter A. Turn on the power switch on the left hand side of the colorimeter. colorimeter. Allow 15 minutes for the instrument to warm up.
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Select the absorbance mode by pressing the MODE button in the lower left until the display reads ABS in the right hand side of the LED display. Turn the dial on the top to the desired wavelength (e.g., 600 nm). Fill the the “blank” cuvette with reverse osmosis osmosis (RO) water, and place it in the rear cuvette holder. Turn the dial on the top of the colorimeter to the word word BLANK position. Close the lid, and press the CAL button. Fill the “sample” cuvette with with RO water, and place it in the front cuvette holder. Rotate the dial to the SAMPLE position. Then close the lid and read the absorbance. If the absorbance reading reading is nonzero, subtract subtract this reading reading from any subsequent readings using that cuvette, or find another cuvette that gives a reading of zero with RO water. Replace the distilled water water in the “sample” cuvette with 2 to 3 mL of sample, and return the the cuvette to the front holder. Close the lid and record record the absorbance reading. Suggestions for using the colorimeter 1. When handling cuvettes, touch only the frosted sides of the cuvette, and wipe any smudges or dust from the clear windows with a Kimwipe before taking absorbance readings. Load the cuvettes into into the colorimeter so that the light passes through the clear windows. windo ws. 2. The colorimeter is sensitive to the position of the cuvette in the light beam. Experiment by taking cuvettes in and out of the holder and jiggling the dial to develop a reproducible technique before important samples are read. 3. Check the calibration using the blank cuvette occasionally. 4. Warm up refrigerated liquids before placing them in a cuvette. Otherwise, gas bubbles may appear in the liquid, or condensation may form on the outside of the cuvette. Either of these results results will affect the absorbance reading. 5. Keep cuvettes out of the sample holders and at room temperature while they are not being read. Otherwise, they will warm up significantly. significantly. Warming can change the absorbance and cause gas bubbles to form.
