Many laboratory protocols require the serial dilution of reagents or compounds. IC50 assays, commonly used to evaluate drug efficacy, and assay development procedures, as well as standard-curve generation, involve the serial dilution of compounds, proteins, or detection agents. These processes can be streamlined by utilizing automated liquid-handling equipment with serial dilution capabilities. Serial dilution processes face two major challenges. The first is error propagation across columns or rows.
With each sequential serial dilution step, transfer inaccuracies lead to less accurate and less precise dispensing. The result is that the highest dilutions will have the most inaccurate results. To compensate for this error possibility, longer mixing times are required, which then increases the time required to perform the serial dilution. These challenges greatly limit the throughput capacity of an automated serial dilution system.
To overcome these challenges, the effects of various mixing parameters of a serial dilution protocol were explored. Velocity11’s (www.velocity11.com) Bravo™ Liquid Handling Platform performed serial dilution with the same pipette head as a full plate dispenser (Figure 1). With the platform’s VWorks™ software, the application allowed the total control of liquid transfer and mixing heights and speeds, which allowed efficient exploration of mixing parameters.
The goals were to determine which parameters had the greatest effect on mixing and to reduce the time required to perform a serial dilution. Serial Dilution Mix Cycles The basic experiment diluted fluorescein across the columns of a 96-well plate, from A1 to A10 (A11 and A12 were blank wells). The starting volume was 300 µL, and 200 µL tips were utilized for the transfer (150 µL, a 1:2 dilution) and mixing steps (190 µL). There are two main components of an accurate and precise serial dilution: the accuracy and precision of the transfer and the efficiency of mixing. Transfers were previously determined to have a precision and accuracy of 99% at this volume; any observed deviations in precision and accuracy were due to error propagation from ineffective mixing.
Two measures were used to evaluate mixing efficiency. The Coefficient of Variance (CV) of each column indicated the precision of the mixing step.
The CV also provided information on the propagation of error across a plate—the CV increased sharply across the plate if mixing was incomplete. The second indication was the accuracy of the transfer. A calibration curve was prepared, and each experimental dilution concentration was plotted against the standard curve to determine the real concentrations in each column. The first experiment varied the number of mixing cycles between 3 and 20. The average precision (averaging CVs for columns 1–10) improved asymptotically as the number of mix cycles increased. Three mixes before each transfer yielded an average CV of 11.8%, while 20 mixes gave a considerably better CV of 1.7%.
The precision in all cases generally worsened as the serial dilution proceeded across the plate; this was expected as the error in the earlier columns propagated with each transfer. The accuracy ratio improved as the number of mix cycles increased. The accuracy ratio is an average of the concentration of the diluted column compared to the previous column—a perfect serial dilution has an accuracy ratio of 1:2.00 across the entire plate. The accuracy ratio of the plate improved with more mix cycles, improving from 1:1.85 to 1:2.01. While the precision and accuracy with 20 mix cycles is close to a perfect serial dilution, the length of time required might be considered impractical.
The 20-mix cycle protocol required 20 minutes per plate, while a three-mix cycle protocol required less than six minutes. Efforts were then focused on the factors that could improve the three-mix cycle protocol to produce accuracy and precision results consistent with the 20-mix cycle protocol. Mix Tip Height The mix tip height was modified in order to determine the effect of distributing the liquid at different locations in the well. As the mix tip height was raised, the average precision improved. At a height of 3 mm from the bottom of the well, the average precision was 3.9%.
The precision worsened as the tip distance from the bottom of the well decreased, reaching a CV of 15% at a height of 0.1 mm. Accuracy tracked with precision, and the higher mix height also improved the accuracy ratio to 1.95. This trend is possibly because the higher dispense height ensures that more of the sample was circulated by the mix cycle.
In a mix roughly in the middle of the well volume, dispensed liquid is forced toward the well bottom while dispensing, and aspirated liquid is pulled from the center of the well. If the mix occurs close to the bottom of the plate, the dispensed liquid is pulled back into the tip during the aspiration. Mixing in the center allows the dispensed liquid to be more evenly distributed in the sample, thus increasing the likelihood of efficient mixing.
Mix Liquid Class Setting The VWorks software controlling the Bravo platform allows the creation of liquid classes, which allows the operator to modify the velocity and acceleration for aspirating, dispensing, and mixing tasks. The original liquid class settings for the mix were 100 µL/s velocity and 500 µL/s2 acceleration. Precision and accuracy improved as the mix velocity increased. This effect plateaus; above 300 µL/s, there is no appreciable improvement in increasing the speed.
The cause of this is likely due to the creation of more turbulent mixing, which in turn distributed the fluorescein dye more quickly throughout the solution. Dynamic Tip Retraction/Extension Finally, the effect of dynamic tip retraction and extension was explored. This function moved the tips deeper into the well during each aspirate step, and retracted them during each dispense step. This allowed a larger volume of the well to be effected by the mix step by adding the movement of the tip into the mix task. There was a marginal improvement (less than 0.5% improvement in CV/accuracy) observed in using this technique.
Additionally, no effect was observed by utilizing another mix standard, which involved aspirating close to the bottom of the well and dispensing near the top of the solution. This mixing method caused no improvement once the other parameters described above had been optimized. These experiments mixed homogenous solutions; there may be an improvement with this technique if the solutions are expected to have different viscosities. Based on these experiments, the parameters that had the largest impact on efficient mixing were (in decreasing order):. Speed of the mixing step. Height of the tip during the mix.
Tip-retraction capabilities To verify this conclusion, the first experiment (varying the number of mix cycles) was repeated with the improved mix parameters. The new parameters provided increased precision and accuracy, and improved the accuracy and precision of the 3-mix cycle operation to a level comparable with the 20-mix cycle operation (Figure 2). More importantly, the new parameters also decreased the time required to run an effective serial dilution protocol from 20 minutes to just under 5 minutes. This has tremendous potential in automating a serial dilution assay and ensuring accurate and precise results.
Part C: UV Experiments Serial Dilutions and Viable Cell Counts The experiment Observing the Effects of Solar Ultraviolet Radiation on Cells shows that when cells are exposed to sunlight all, some, or none of them may be killed. Many experimental questions can be answered with qualitative answers like 'all, some, or none.' Other questions may require quantitative answers. For example, in the next experiment you will use the sensitive yeast strain to measure the intensity of solar UV radiation by measuring the fraction of cells exposed that survive. To get quantitative answers about yeast survival you must put a known numbers of viable (living) cells onto the agar plates and then count the number that remain after being exposed. You can determine the number of viable cells by counting the colonies that grow up on the agar growth medium in a Petri plate by assuming that each colony grows from a single viable cell. This is usually a reasonable assumption.
Experiment: In the experiment that follows you will learn how to measure the number of viable cells on a Petri plate. You will be able to use this procedure whenever you need to measure the number of cells that survive an exposure to radiation or some other treatment. First you will estimate the number of cells in a liquid suspension in order to plate a reasonable number of cells. For this you will use one of the most sophisticated and sensitive optical instruments in existence, the human eye. With surprisingly little practice you can learn to estimate the number of cells in a suspension by just looking at it.
You can estimate cell density because of your eyes' fairly sharp threshold for observing turbidity (cloudiness). When viewed in a standard 13 100 mm glass tube, yeast suspensions of less than about 1 million cells per mL are not visibly turbid. Above this threshold density, the suspension is cloudy. When you adjust the number of cells in a suspension until just barely visible, you obtain a suspension of known density (approximately 1 106 cells/ml). When you have a suspension that contains approximately 1 106 cells/ml, you will dilute it to get the right concentration for plating. You will make the dilutions in known steps so you can calculate the number of cells in each dilution tube.
This procedure helps you plate a countable number of colonies.
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Pre Lab: Serial Dilutions Practice Exercise If students are not familiar with operating pipettes, the Pre Lab Serial Dilutions practice exercise is recommended. The exercise will allow students to learn how to pipette, dilute samples and calculate the concentration of their samples.
The use of methylene blue gives them a visual model of how samples become less concentrated with each dilution step. This activity was adapted from An Introduction of Biotechnology developed by the Mathematics and Science Education Center, University of Missouri- St. A student answer sheet and teacher Power Point are attached for use during this lab. The Pre Lab Exercise can be completed in forty five minutes. Materials Needed Pre Lab Serial Dilutions Equipment and materials for students Equipment and notes Quantity per group (recommend 2-4) □ Test Tube Rack 1 □ Test Tubes 4 □ Flask (recommend 1000mL) 1 □ Pipette Pump (10 mL) 1 □ Pipette (10 mL) 1 □ Methylene Blue (preferably in a dropping bottle) 6 drops □ Water 1 Full Flask □ Lab Tape 2 inches □ Lab Coat or Apron 1 per person □ Safety Glasses 1 per person □ Safety Gloves 1 pair per person □ Calculator (Optional) 1 Preparation Instructions Set up the test tube racks with four test tubes per group. Place the other equipment and supplies at the student work stations or in a central location where they can be located by group members and taken to their work stations.
Class Demonstration/Pre-activities If your students are not familiar with how to use a pipette and pump, review the proper way to attach the pipette and pump and how to hold the equipment and how to measure accurately with the pipette. Also, review how to mix the solutions in the tube by using the pipette to draw in the contents of the tube and then releasing back down.
Repeat that step two times for a full mix. Pre Lab Definitions Use the to review the pre lab definitions that the students have on their lab sheet. Serial: In a series, order or interval. Measured steps. Dilution: Water Down. Pipette: “Little pipe” for moving and measuring liquids.
Serial dilutions reduce the concentration of a sample in small steps or fractions. Conducting the Lab The can be used to reveal instructions to the students during the lab.
The teacher may wish to go step by step, making sure that all groups have completed a step before moving on to the next one. This will allow the teacher to view the student’s pipetting technique and correct any problems that arise. Students should wear protective aprons or lab coats, gloves and safety glasses due to the use of methylene blue during this lab.
We propose a simple technique for bacterial and yeast cfu estimations from diverse samples with no prior idea of viable counts, designated as single plate-serial dilution spotting (SP-SDS) with the prime recommendation of sample anchoring (10 0 stocks). For pure cultures, serial dilutions were prepared from 0.1 OD (10 0) stock and 20 μl aliquots of six dilutions (10 1–10 6) were applied as 10–15 micro-drops in six sectors over agar-gelled medium in 9-cm plates. For liquid samples 10 0–10 5 dilutions, and for colloidal suspensions and solid samples (10% w/v), 10 1–10 6 dilutions were used. Following incubation, at least one dilution level yielded 6–60 cfu per sector comparable to the standard method involving 100 μl samples. Tested on diverse bacteria, composite samples and Saccharomyces cerevisiae, SP-SDS offered wider applicability over alternative methods like drop-plating and track-dilution for cfu estimation, single colony isolation and culture purity testing, particularly suiting low resource settings.
1. Introduction Estimation of colony forming units (cfu) through serial dilution plating on a nutrient medium forms the most widely accepted method for monitoring cultivable bacteria and yeasts in different spheres of microbiology,. Cultivation-based methods being simple to practice, command enormous significance and applications in bacteriology.
This holds good in spite of the emergence of molecular techniques such as fluorescent in situ hybridization, real-time quantitative PCR, flow cytometry, etc., which although provide a precise account of metabolically active cells, demand much expertise and resources. Further, cfu-based techniques provide information on the most abundant populations among the cultivable community,. Viable colony counts also form essential tools in biotechnology such as gene cloning, surveillance of genetically modified organisms, assessing bioremediation effects, testing novel anti-microbials, etc. Besides serving as standards during molecular investigations.
Spread-plating and pour-plating form the standard approaches for bacterial and yeast cfu estimations,. Spread-plating offers several advantages over pour-plating such as more flexibility in handling, less interfering effects on temperature sensitive organisms, the avoidance of aerobic organisms getting trapped inside agar medium, the surface enumeration of cfu and the easy selection of distinct colony types,. Here, the bacterial sample is applied over agar-gelled nutrient medium with the help of a glass, plastic or steel spreader where the spreader is generally considered a mere tool to distribute the inoculum over the agar surface,. We have documented that the inoculum-spreader employed during standard spread-plating could impart significant injury to bacterial cells and affect the cfu depending on the extent of its usage on the agar surface. This was demonstrated in comparison with the alternate approach that did not involve the use of spreader, namely, spotting- and- tilt- spreading (SATS). Any spreader movement on agar surface subsequent to the exhaustion of free moisture proved detrimental to the bacterial cells further influenced by the operator practices and moisture levels in the medium.
The physical impaction effects on vegetative cells varied between different organisms governed by the cell characteristics of the bacterium with Gram-negative organisms being more vulnerable than Gram-positive bacteria, cocci less susceptible than rods and more risk to larger cells than smaller cells. The physical impaction effect also applied to the supposedly hardy spores of Bacillus spp. Which seemed comparable to glass globules that crumble under physical pressure. Thus, the spreader-independent SATS approach proved to be a simpler and safer alternative to spread-plating for bacterial cfu estimations with several other advantages,.
Generally 25–250 or 30–300 colonies per agar plate (100 μl sample) are prescribed as the acceptable cfu for accurate counting,. When there is no clear indication of the dilution level that yields this cfu range, several plates representing different dilutions and replications need to be employed leading to considerable wastage of time, manpower and material resources,. This applies invariably to pure bacterial cultures, water, food, soil and various environmental and biotechnological specimens. As we found that inoculum-spreader was wholly dispensable, accommodating multiple dilutions in a plate was considered. Similar attempts in the past included drop-plating, track-dilution and drop-spotting with digital imaging, but these studies used pure bacterial cultures that yielded confined colony growths.
The situation is different when the samples involve fast growing organisms, mixture of different bacteria varying in growth rates or colony characteristics, and with food and environmental samples. The present studies were undertaken to optimize a simple and resource saving method for bacterial cfu estimations that allows the accommodation of multiple dilutions in a plate and to test the feasibility of the technique across diverse samples including pure bacterial and yeast cultures and composite samples. Bacterial and yeast cultures and composite samples Pure cultures of bacteria belonging to different phylogenetic groups varying in Gram reaction, cell characteristics and sporulation potential were used towards optimizing the single plate-serial dilution spotting (SP-SDS) technique employing spotting- and- tilt- spreading (SATS), as the standard procedure. The organisms included Enterobacter cloacae, Escherichia coli, Acinetobacter junii (Proteobacteria), Bacillus pumilus, Bacillus subtilis, Bacillus thuringiensis, Staphylococcus epidermidis, Staphylococcus haemolyticus (Firmicutes) and Microbacterium esteraromaticum (Actinobacterium) described elsewhere. Cloacae was used as the primary candidate for protocol optimization followed by B. One strain of ascosporogeneous wine yeast ( Saccharomyces cerevisiae) was used in this study employing potato dextrose agar (PDA). Different composite samples representing public health, food, environmental, agricultural, clinical and biotechnological settings described below were also tested for bacterial or yeast cfu.
Additionally, an endophytic bacterial strain of Pseudomonas aeruginosa from banana that could be monitored distinctly from other organisms on cetrimide- nalidixic acid- agar (CNA) selective medium was used as a representative of clinical specimens and genetically modified organisms. Unless mentioned differently, overnight nutrient agar (NA)/nutrient broth (NB) derived (18–24 h) cultures were used in all studies involving pure bacterial cultures except for spores. Nutrient media Nutrient agar sourced from M/s HiMedia Biosciences (Mumbai, India) formed the standard bacteriological medium while the other media formulations mentioned later were employed for specific organisms/samples and also to test the applicability across different media. Unless mentioned differently, NA/fresh PDA prepared in pre-sterilized disposable Petri-dishes on the same day about 2 h post-pouring (referred to as fresh plates) or that prepared on the previous day and incubated overnight at 37 °C after sealing in polypropylene (PP) bags were used in all trials. The nutrient plates used in a specific trial belonged to the same batch of preparation unless mentioned differently. SP-SDS and SATS procedures For pure bacterial and yeast cultures, a uniform cell suspension was prepared by dispersing the overnight colony growths from agar plates, or NB culture after one spin-wash in sterile water in the case of Bacillus spp.
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After allowing any cell clumps to settle down, the clear upper part was transferred to a fresh tube. The optical density (OD) was determined at 600 nm employing a 1:10 diluted stock in a uv/vis spectrophotometer (Genesis 10 UV, Thermo Scientific, MA, USA) based on which the ‘anchored stock’ of 0.1 OD (10 0) was prepared. Decimal serial dilutions (100–1000 μl) of 10 1–10 6 or 10 7 were prepared from the 10 0 stock in 1.5 ml tubes with 4–5 repeated flushing and changing of tips (see movie: ). For preparing the stock and serial dilutions, filter-sterilized distilled water (FDW) aliquoted and stored at −20 °C was preferred unless the water was freshly autoclaved.
This was essential to avoid the chances of any hardy autoclaving defying spores multiplying during the post-autoclaving storage. Spore preparations and dilutions were made from 7-day old NA plates in 50% ethanol as described elsewhere to avoid their germination. For water and clear liquid specimens, the direct sample formed the 10 0 anchored stocks. Thick and colloidal suspensions such as milk and fruit juice were used directly or after adjusting OD600 nm to 1.0 or 10 while for solid specimens (food, soil) a suspension prepared in water at 1.0 g sample per 10 ml formed the 10 0 stock. In this study, our emphasis was on cfu enumeration technique rather than sampling methods for which the accepted standard procedures prescribed were to be adhered (e.g., ).
To execute SP-SDS, the reverse of the 9-cm Petri-dishes containing surface dry agar media were drawn to six sectors with the marking of first and last dilution sectors for clear identification. Using a calibrated micropipette, 20 μl aliquots from selected six dilutions were applied as 10–12 micro-drops in these demarcated areas.
During sample spotting, the same tip was used starting with the lowest dilution. Care was exercised to avoid tip marks on the medium during sample application not to mistake them for cfu. The sterility of the diluent was ensured by spotting 20 μl at the bottom part of the plate. The plates were exposed in the laminar air-flow (LAF) cabinet for the droplets to dry off (8–10 min for fresh plates and 3–4 min for pre-prepared surface-dry plates), sealed in polypropylene (PP) covers and incubated inverted at 28–37 °C as required for specific organisms. For SATS, 100 μl of different dilutions were applied as 20–25 micro-drops per plate and spread on agar surface by mere tilting or gentle twirling of plate followed by surface drying (5–6 min) in the LAF,.
Cfu enumeration was done after 18–48 h with the marking of colonies on the reverse of the plate. An illustration of single plate-serial dilution spotting (SP-SDS) technique for pure bacterial / yeast cultures and composite samples. The colony development pattern at different dilutions in SP-SDS was recorded as spot growth, too many to count (tmtc) or countable/acceptable (6–60 range). After recording the dilution level yielding acceptable colonies and the cfu per sector, cfu per 100 μl was worked out as n × 5 ( n = colonies in 20 μl sample applied area).
The cfu ml −1 of the 10 0 stock was arrived at as the product of n × 5 × 10 d+1 (d = dilution level yielding the countable colonies). Assessing intra- and inter-plate variations in SP-SDS employing E. Cloacae and mix inoculum To get an estimate of the possible sector to sector variations in a plate or inter-plate variations during SP-SDS, E. Cloacae serial dilutions of 10 4 and 10 5 were spotted (20 μl) in three sectors each in ten fresh NA plates of which 10 4 yielded tmtc and 10 5 countable colonies.
The cfu per sector (average of three sectors at 10 5), standard deviation (SD) and coefficient of variation (CV) were worked out for each plate individually. A similar experiment was undertaken employing the mixed inoculum of five organisms ( E. Thuringiensis, S. Epidermidis and M. Esteraromaticum) which were pooled in equal proportions employing the dilution levels that yielded the acceptable cfu (30–300 per 100 μl). Assessing the number of replications needed for comparable cfu estimates in SP-SDS and SATS SP-SDS was undertaken in comparison with SATS using E. Cloacae 10 5 dilution.
SATS involved 100 μl sample applied in 12 NA plates while in SP-SDS, 10 5 dilution was applied in six sectors in 12 plates. Colony counts were made adopting one sector per plate sequentially representing the six sectors across 12 plates in SP-SDS and the cfu/100 μl was recorded in both methods. The mean, SD and CV were worked out sequentially for 2–12 replications. Further, the data were tested for significance through single factor ANOVA considering two to 12 replications sequentially. ANOVA between SATS and SP-SDS was also done considering the average cfu counts from the six sectors in the 12 SP-SDS plates.
The experiment was repeated employing a composite sample comprising of E. Thuringiensis, S.
Epidermidis and M. Esteraromaticum prepared as above but in irrigation grade tank water with a prior SP-SDS assessment of cfu to fix the appropriate dilution level. Testing SP-SDS versus SATS on additional pure cultures and composite samples The applicability of SP-SDS was tested employing pure cultures of different bacteria.
This included E. Aeruginosa, A. Thuringiensis, S. Epidermidis, S. Haemolyticus and M. Esteraromaticum employing NA at 30 °C except for E.
Coli for which trypticase soy agar (TSA; 37 °C) was used. Employing d1–d2 source cultures, 0.1 OD (10 0) stocks were prepared in FDW followed by the preparation and usage of six decimal serial dilutions. A similar experiment was undertaken with the yeast strain on PDA. The composite samples included irrigation-grade tank water, milk, ground mixed-vegetables and a soil sample.
For pure bacterial cultures, the tested dilutions included 10 1–10 6, for clear water 10 0–10 5, for milk, ground vegetables and soil, 10 1–10 6 avoiding the particulate 10 0. Cfu enumeration was done manually after 18–48 h and beyond as needed depending on the organism/sample. Based on the information of the appropriate decimal dilution that yielded 30–300 cfu per 100 μl sample, SP-SDS was undertaken in comparison with SATS employing four replicate plates for SATS and adopting cfu from first four of the six sectors in an SP-SDS plate for statistical analyses.
Two independent serial dilutions were prepared each applied in duplicate SATS plates or three SP-SDS sectors each. The cfu counts were translated to cfu ml −1 of 10 0 stock and analyzed for significance employing single factor ANOVA (Microsoft Excel 2010) after logarithmic transformation.
Testing SP-SDS approach across other media and NA plates of different batches SP-SDS approach was tested across other media including Luria Bertani agar for E. Coli, plate count agar, brain heart infusion agar, Muller Hinton agar and MacConkey agar for E. Cloacae and irrigation grade water.
CNA medium was tested employing P. The media formulations were sourced from M/s HiMedia Biosciences, Mumbai. Based on the earlier documentations that the quantity of medium per plate, the age of plates after the preparation and the pre-treatments given to the plates did not alter the cfu estimates in SATS, fresh plates with 15, 20 or 30 ml NA were tested in SP-SDS for the time needed for droplet drying and the cfu after applying the 10 5 dilutions of E. Cloacae and B. Further, 20 ml NA plates prepared on the same day or that prepared 1–7 days before and the plates given a 37 °C pre-warming treatment were tried in SP-SDS wherein fresh 20 ml NA plates served as control. The experiments were repeated wherein B.
Pumilus culture was employed at a non-decimal dilution (1:3 of 10 4) to get more acceptable cfu range (100 per 100 μl) as in the earlier study. Testing SP-SDS methodology employing multi-well plates SP-SDS methodology was tested with E. Cloacae and B.
Pumilus using 96 cavity (500 μl) autoclavable polypropylene assay plates (Cat. P.96-450R-C; Genaxy Scientific Pvt., Ltd., Solan, India) for serial dilutions adopting 40–400 μl or 50–500 μl decimal dilutions (10 1–10 6).
As controls 100–1000 μl and 40–400 μl dilutions in 1.5 ml microfuge tubes were employed. Additionally, ELISA plates (Greiner Bio-One GmbH, Germany) were tried which accommodated 200 μl sample per well employing 20–200 μl dilution series. Testing of SP-SDS methodology for microbiological and biotechnological samples Different samples representing biotechnology, agriculture, medicine, food microbiology, environmental microbiology and applied microbiology where there was no clear idea about the prevalent bacterial or yeast cfu in the sample were tested through the SP-SDS approach. The preferred dilutions from the anchored stocks included 10 0–10 5 or 10 1–10 6 for liquid samples, and 10 1–10 6 for solid samples avoiding the particulate 10 0. Further, SP-SDS was tried for parallel testing of two or multiple samples in a plate. This included testing the effect due to different diluents on E.
Cloacae where the 10 0 stock in FDW was taken through serial dilution in saline (NaCl 9 g l −1), phosphate buffered saline (PBS), peptone–water (10 g l −1 peptone and 5 g l −1 NaCl; pH 7.2), peptone-salt (1 g l −1 each peptone and NaCl; pH 7.0; ) or nutrient broth (pH 7.4) employing FDW as control. In another trial, E. Cloacae dilutions prepared in FDW and peptone salt was monitored with SP–SDS after static incubation over 5 h at 20 min intervals during the initial one hour and hourly thereafter employing the decimal dilutions 10 3–10 8. Further experimental details are provided under Results and Discussion. Statistical analysis For direct comparisons within SP-SDS trials, the mean colony counts per sector in a plate and for comparisons between SP-SDS and SATS techniques, cfu per 100 μl samples were used for statistical analysis. The mean, SD and CV were employed for direct comparisons estimated with the Σ function in Microsoft Excel 2010.
In the trial comparing SP-SDS versus SATS, the significance was tested through single factor ANOVA or Student’s t-test using the Data Analysis Tool of Microsoft Excel 2010 after logarithmic transformation of cfu for the 10 0 stocks. Unless mentioned differently, four replications were employed for comparing SP-SDS versus SATS. Preliminary SP-SDS trials In the initial trial employing E. Cloacae, the first three serial dilutions (10 1–10 3) showed spot growth, 10 4 displayed tmtc and 10 5 yielded well delineated colonies in the acceptable range (A). The plates applied with the irrigation-grade tank-water exhibited cluster of diverse colony types at 10 0–10 2 including some spreaders and at 10 3 countable colonies (B). Thus, at least one dilution in a plate yielded cfu in the acceptable range ensuring the success of the trial. Cloacae displayed full colony emergence by day-1 itself whereas colony development continued for 2–4 days for irrigation water.
Marking the initially formed colonies on the reverse of plates helped in identifying late emerging ones and discriminating the fast-growing or spreading colony types. Vortexed and non-vortexed samples of E. Cloacae showed similar cfu for both the treatments (data not shown) indicating that vortexing during serial dilutions was not a necessity for homogeneous suspensions but the same did not impart any adverse effect.
Assessing intra- and inter-plate variations during SP-SDS E. Clip studio paint serial number generator. Cloacae 10 4 dilution showed tmtc while 10 5 dilution exhibited cfu in the range of 27–41 per sector with the mean sector cfu of 31.3–38.3 across 10 different plates. No significant plate to plate cfu variations were observed in the trials employing E. Cloacae ( P = 0.553) and the mixed inoculum ( P = 0.0673).
The cfu per sector for the mixed inoculum varied from 19 to 34 across 10 plates and the mean cfu per sector in a plate ranged from 24.0 to 29.8. Assessment of the number of replications needed for comparable cfu estimates in SP-SDS and SATS The ANOVA results with E. Cloacae employing 2–12 replications indicated statistically comparable cfu for SATS and SP-SDS ( P 0.05 in all instances) starting with two replications (A). The same appeared true for the mixed inoculum prepared in irrigation-grade water (B).
Based on these results and the observations from the subsequent trials, use of four replications for SP-SDS was fixed to give similar cfu as in SATS. No definite advantage of using 4 replications was observed based on mean, SD and CV for E.
Cloacae and for the mixed inoculum. Testing SP-SDS versus SATS on additional organisms and composite samples Adopting SP-SDS with pure bacterial cultures, most of the colony development occurred within 18–24 h and within 2 days for slow growing organisms like M. Esteraromaticum and A.
The 10 0–10 2 dilutions from the anchored stocks often displayed spot growth, 10 3–10 4 tmtc and 10 3–10 6 isolated colonies depending on the organism apparently governed by the cell size as per the earlier report. For instance, B.
Thuringiensis with large rods and S. Epidermidis with large cocci showed countable cfu at 10 4 dilution; E. Aeruginosa, B. Subtilis and M.
Esteraromaticum with medium-size rods or S. Haemolyticus with smaller cocci at 10 5, and A. Junii with very small cells at 10 6. This was confirmed in repeat trials when M. Esteraromaticum and A. Junii showed some variations with the acceptable cfu at 10 5 or 10 6 in some trials. As for Bacillus spores, the dilution level for acceptable cfu also varied with the organism (10 3–10 4) depending on spore size as documented earlier,.
This applied to pure yeast culture too (10 3). The composite samples- mixed inoculum, water, milk, food articles and soil samples- also showed spot growth, tmtc or acceptable cfu depending on the dilution level. Nearly 80–90% colonies emerged within 1–2 days but it required 3–4 days to allow most colonies to develop. Spreaders were occasionally encountered for water, milk and soil samples, particularly with incubation beyond 3–4 days.
This applied equally to SATS, spread-plating and pour-plating (Thomas, unpublished results). BANOVA after logarithmic transformation; NS, not significant.
SP-SDS worked satisfactorily for all test organisms and composite samples giving acceptable cfu at least in one of the six dilutions from the anchored stocks in a plate. Now, comparing SP-SDS with SATS, the mean cfu recorded for the 10 0 stocks of pure bacterial cultures or spores in different organisms, yeast and in composite samples over four replications appeared statistically on par.
The SD recorded in SP-SDS was higher in all instances (average cfu-SD for the 19 samples listed in, in SP-SDS and SATS, 1.44 × 10 7 and 9.3 × 10 6, respectively) but the means from four replications appeared close to each other. Thus, SP-SDS with the anchored stocks offered assured and comparable results to SATS with mere four nutrient plates as against 24 plates needed for a similar testing through conventional plating approaches. For organisms such as B. Thuringiensis and A. Junii, the cfu registered for the selected decimal dilution was close to the lower acceptable range while the previous decimal dilution level showed tmtc. This was influenced by the cell size, which is a characteristic feature of the organism.
In such instances, the adoption of non-decimal dilutions ( e.g. 1:3 of 10 4 for B. Pumilus and B. Subtilis) were tried for clearer results which showed identical cfu in SATS and SP-SDS (data not shown). Comparison of SP-SDS with alternate resource saving approaches A comparison of SP-SDS with 6 × 6 drop-plating and track-dilution indicated that the former be advantageous over the two other methods in several respects (; ). The 6 × 6 drop-plating was fine for slow growing pure cultures but not for fast growing colony types and for composite samples that bore organisms differing in colony growth rates. SP-SDS worked well for all samples including pure cultures, spores and composite/environmental samples.
This also applied to track dilution but for the high cost of square plates. Testing SP-SDS method across other media and NA plates of different batches SP-SDS approach with decimal dilutions worked well across different media yielding well delineated colonies for pure cultures of most of the organisms. Subtilis and B. Thuringiensis showed tendency for fast or spreading colony development. Testing NA plates with different amounts of medium, E.
Cloacae showed identical cfu in plates with 15, 20 or 30 ml fresh medium (50.8, 51.0 and 53.3 per sector at 10 5 dilution, respectively; NS). The same appeared true for B. Pumilus (11, 11.5 and 11 cfu per sector, respectively, for 10 5 stock and 28.6, 27.0 and 24.0, respectively in the trial employing 1:3 dilution of 10 4 stock; NS).
This indicated considerable saving of media resources with the use of 15–20 ml medium per plate. Besides, the time needed for the droplet drying was considerably shortened with the reduction in the amount of medium per plate (4, 6 and 10 min, respectively for 15, 20 and 30 ml fresh NA plates). The colonies in 15–20 ml appeared smaller and stayed confined for longer time than in 30 ml plates. One stipulation with SP-SDS was the proper drying of droplets in the LAF cabinet.
Testing fresh 20 ml NA plates versus media prepared 1–7 days before (sealed in PP bags), or refrigerated for a month, the cfu estimates for E. Cloacae were unaltered between them, but the older plates offered considerable saving of time towards sample drying.
The time for droplet drying was governed by the free moisture content in the medium. For instance, freshly poured NA plates (20 ml) used within 1 h required about 6–8 min which was reduced to 4–5 min by 2 h after pouring and to about 1–2 min if used after one or a few days after preparation. Pre-warming the plates at 37 °C was not a necessity nor offered any advantage but often caused water condensation which ought to be removed before using the plates. Variable durations of open plate incubation in the LAF did not alter the cfu for up to 60 min as observed with E. Cloacae and B. Pumilus cultures (; P 0.05 in both instances) allowing flexibility in the operations. Thus, SP-SDS technique worked fine with freshly prepared plates, previously prepared refrigerated or ambient stored plates (sealed in PP covers) and even plates with partial dehydration, and also worked well with varying amounts of medium per plate as per the observations employing E.
Cloacae similar to the findings with SATS technique. SP-SDS methodology with multi-well plates Testing SP-SDS method with the use of 96 cavity assay plates for serial dilutions showed that 40–400 μl dilutions was feasible but not 50–500 μl series due to the chances of inoculum mixing between adjoining wells. Comparing 40–400 μl dilution series in assay plates versus 40–400 μl or 100–1000 μl dilution in 1.5 ml microfuge tubes showed similar cfu in the three treatments for E. Cloacae (39.0 ± 3.93, 44.5 ± 5.54 and 36.5 ± 2.5 cfu per sector, respectively; P = 0.108). The corresponding figures for B. Pumilus were 11.7 ± 5.60, 11.7 ± 2.21 and 14.0 ± 4.54 ( P = 0.711). ELISA plates were not preferred as they could accommodate only smaller volume (200 μl) besides their high cost and non-feasibility for reuse unlike the autoclavable assay plates.
Demonstrating the applications of SP-SDS in microbiology and biotechnology SP-SDS method worked well for various samples tested where there was no prior idea of the dilution level that would yield countable cfu giving acceptable colony counts for at least one of the six dilutions contrary to the six plates required in SATS, spread-plating, pour-plating or spiral plating to accommodate same number of dilutions. Besides pure and mix cultures, these included environmental samples, probiotic and agricultural bio-formulations, cultures of different organisms post antibiotic challenge and different food products. For food samples and other instances where the viable counts or the microbial composition would change with time or storage, SP-SDS formed an ideal tool for cfu assessment.
Use of 4–6 replicate plates is recommended for testing such items that could not be stored or retested. The utility of SP-SDS technique was noteworthy while testing broth cultures in different stages of growth or an organism grown under different conditions, testing the effect due to antibiotics and other antimicrobials on single organisms or mix-cultures where the extent of cfu reduction varied depending on the organism and the chemical employed. Water and soil samples introduced with clinically significant P.
Aeruginosa ( Pau) could be specifically monitored for Pau on CNA selective medium with parallel testing on NA to assess its load and the interactive or inhibitory effects on other microflora. The distinct green-tinge of fluorescent Pau colonies allowed their clear identification on NA yielding similar counts on CNA medium Thomas and Sekhar, unpublished results. The SP-SDS method also worked satisfactorily for market lots of active dry yeast yielding delineated colonies on PDA. While pure yeast culture displayed more or less uniform colony emergence, the market lots showed colony development spanned over 2–3 days. Experimental sample/specimen Anchored stock Dilutions tested Dilution yielding cfu CFU range/sector Av. Significance of SP-SDS methodology and further optimizations The hall mark of SP-SDS was ensuring acceptable cfu at one of the decimal dilutions thereby safeguarding against the failure of cfu assessing trials. SP-SDS was particularly useful for side by side testing of two or multiple samples.
For instance, we were eager to determine the most appropriate diluent without adverse or contributory effects on cfu due to bacterial cell lysis or multiplication during the SP-SDS procedure. Testing six different diluents for E. Cloacae which could be done by accommodating the six treatments in a single plate for their direct comparisons, the maximum cfu was recorded for peptone-salt followed by FDW, saline, PBS and peptone-water (on par) while NB registered a notably lower cfu (A). To ascertain the lower colony counts in NB, we further tested NB in comparison with FDW and peptone-salt (PS) as controls on E.
Cloacae and B. This again indicated a lower cfu with NB in E. Cloacae (46.8, 39.6 and 37.0 cfu per sector for FDW, PS and NB, respectively) as well as for B. Pumilus (12.0, 8.8 and 8.3, respectively, using 10 5 dilution, and 34.2, 31.2 and 18.5, respectively, for 1:3 dilution of 10 4).
The undesirable NB effect appeared to arise from the inhibition to cell germination due to higher nutrient levels at the sample dried spots. Further, testing the growth of the two organisms in 1.0× and 1.25× NB with overnight shake incubation indicated that the growth was not enhanced but rather reduced at higher NB level in both E. Cloacae (OD600 nm of 1.696 and 1.346, respectively; P = 0.028) and B. Pumilus (1.212 and 1.033, respectively; P = 0.001). Thus, it appeared that NB was not a preferred diluent while FDW and PS appeared fine.
SP-SDS applications in microbiology and biotechnology: Feasibility of accommodating up to six replications of a selected dilution in a single plate in SP-SDS (A1) in comparison with one replication per plate in SATS (A2); Parallel testing of six different organisms (left to right from top: Bacillus pumilus, B. Subtilis, Escherichia coli, Enterobacter cloacae, Staphylococcus epidermidis and Microbacterium esteraromaticum) on nutrient agar (B1) versus Luria Bertani agar (B2) with E. Coli showing delayed growth on NA at 30 °C; SP-SDS showing culture admixture where the contaminant appears as large colonies at 10 5 and 10 6 dilutions (C).
Serial Dilution Lab Report
Most of the environmental specimens employed in this study showed acceptable cfu within the first four dilutions. Applying four dilutions (10 1–10 4) in four sectors per plate allowed more area per sample accommodating diverse and even spreading colony types. No differences in cfu per sector were observed if four or six sectors were prepared in a 9-cm plate as observed with E. Cloacae (42.6 ± 7.55 and 47.6 ± 6.91, respectively) and B.
Pumilus (13.3 ± 3.32 and 10.8 ± 3.48, respectively). When diversity analysis was the objective, running a pre-trial with SP-SDS helped in identifying the preferred dilution based on which SATS trials could be set up to cover low abundant types. Although the usage of known sample weight per unit volume for solid food articles or the use of direct samples for liquid specimens as starting stock is a common practice in microbiology, none of the publications specifically emphasize the need for ‘sample anchoring’ as a standard practice. Publications addressing cfu monitoring in pure cultures often use serial dilutions of bacterial suspensions or broths and report final growth assessments based on cfu and OD rather than anchoring the OD initially. The concept of accommodating multiple serial dilutions in a nutrient plate is also in vogue in bacteriology,. The significant aspects of this study have been the prescription of sample anchoring to a specific and reference base (10 0 stocks) as a standard practice at the start of the trial plus the accommodating multiple dilutions in a plate. Anchoring the specimens ensured that at least one dilution level yielded acceptable colony counts in a plate and that the experiment would not fail wholly in the absence of which some trials overshot the acceptable cfu level in a plate.
With the identification that the cfu ml −1 in an organism at a particular OD is governed by cell or spore size, we are now able to set up SATS trials with most organisms at the dilutions mentioned in. In some instances, the non-decimal dilutions were needed to obtain a higher cfu (100 per 100 μl) in critical comparative trials as documented earlier,. It was essential that a relatively thin suspension of 0.1–0.5 OD be used for the OD estimation for precision. For colloidal and thick suspensions, such as milk and juices, the original specimen could be employed directly or after adjusting the OD to a desired level. For instance, milk showed variations in OD600 nm from 150 to over 200 whether it was full cream, toned or skimmed and with brands.
Sample anchoring held good also for other modes of cfu estimation such as SATS, spread-plating or pour-plating. Cfu ml −1 in an organism showed some variations with the source culture medium, age of culture or the way a culture was grown which in turn was attributable to differences in cell size or factors such as cell debris or pigments that alter the OD. For instance, in the results presented herein, E. Cloacae derived from spot-growths showed cfu in the range of 40–50 per sector while that derived from isolated single colonies with larger cells yielded cfu in the 30–40 range. SP-SDS accommodated all such situations with uncertain initial cfu.
The major time investment during cfu estimations was preparing the dilution series which applied equally to SP-SDS, SATS, spread-plating and pour-plating. We are not addressing sampling procedures in this study for which the accepted standard procedures prescribed such as International Commission for the Microbiological Specifications of Foods (ICMSF) or International Organization for Standards (ISO) are to be adhered. Refining the SP-SDS approach further, up to eight dilutions per plate, more amount of sample (25–50 μl) per sector or more area per dilution as for environmental samples and food articles could be accommodated with the use of 10 cm diameter or 12 × 12 cm square plates.
It was important that the decimal dilutions show a clear reduction in cfu with dilution series and that the extinction point (no colonies) is attained within the 10 5–10 7 dilution in the case of pure cultures absence of which indicated improper serial dilution. This was often noticed when tip-flushing and tip-change during onward serial dilution were not adhered to. This also occurred due to the presence of contaminants in the diluent which could occur due to improper sterilization or their accidental introduction during sample handling reinforcing the need for testing the diluent in each plate. Use of FDW is prescribed as the standard diluent without much adverse effects of cell lysis or bacterial multiplication during the course of SP-SDS procedure. It is not proper to use previously prepared and stored stock cultures or dilutions as the organisms display microaerophlic growth even under refrigeration. As a step to automation, it is possible to capture the plate images and effect the colony counts later on.
Thus, SP-SDS appeared advantageous and applicable across different spheres of microbiology and biotechnology for samples of uncertain cfu and under low resource settings. Further, when there is a clear idea of the dilution level for acceptable cfu or for critical comparative trials, we still adopt SATS. 4. Conclusions SP-SDS where six different dilutions of a bacterial suspension or test sample (20 μl) is spotted as micro-drops across a 9-cm plate agar-surface represents a simple, efficient and resource-saving technique for bacterial cfu estimations when there is no clear idea about the initial cfu or the dilution at which countable colonies could be expected.
Sample anchoring (use of 10 0 stock) which in the case of pure bacterial cultures formed the 0.1 OD stock, the original suspension for water and other liquid samples, and 10% (w/v) sample for food and soil specimens, followed by the application of decimal serial dilutions in sterile distilled water ensured that at least one of the dilutions yielded countable colonies in the acceptable range in each nutrient plate. SP-SDS with four replications suited diverse samples including pure bacterial and yeast cultures, spores, mix-bacterial inoculum, food, clinical, environmental and other biotechnological samples giving similar cfu estimates as the standard SATS approach employing 100 μl samples per plate. Besides cfu enumeration, SP-SDS enabled single colony selection and culture purity confirmation.
In vitro evolution of RNA molecules requires a method for executing many consecutive serial dilutions. To solve this problem, a microfluidic circuit has been fabricated in a three-layer glass-PDMS-glass device. The 400-nL serial dilution circuit contains five integrated membrane valves: three two-way valves arranged in a loop to drive cyclic mixing of the diluent and carryover, and two bus valves to control fluidic access to the circuit through input and output channels.
By varying the valve placement in the circuit, carryover fractions from 0.04 to 0.2 were obtained. Each dilution process, which is comprised of a diluent flush cycle followed by a mixing cycle, is carried out with no pipeting, and a sample volume of 400 nL is sufficient for conducting an arbitrary number of serial dilutions. Mixing is precisely controlled by changing the cyclic pumping rate, with a minimum mixing time of 22 s. This microfluidic circuit is generally applicable for integrating automated serial dilution and sample preparation in almost any microfluidic architecture. INTRODUCTION Serial dilution is among the most fundamental and widely practiced laboratory techniques, with applications ranging from measuring detector response, to determining kinetic rate constants, to culturing cells. Serial dilution is particularly important in directed evolution experiments in which a population of RNA molecules is made to undergo repeated rounds of selective amplification.
In order to evolve molecules with interesting properties, it is necessary to propagate the population of RNAs through many logs of selective growth. This is accomplished by serially diluting an aliquot of the reaction mixture into fresh growth medium at regular intervals. Performing serial dilutions by manual pipeting is a mundane and time-consuming task that has limited the execution of highly longitudinal experiments in molecular evolution. Microfluidic technology presents a practical solution to this problem by automating the fluid handling associated with serial dilution.
The core strengths of microfluidic technology are integration, high throughput, and low-volume handling. Microfluidic analogs outperform conventional instrumentation with regard to speed, throughput, and reagent consumption by an order of magnitude or more, and allow integration of sample preparation and analysis in a single device., Precise manipulation of fluids in these devices is achieved by electrokinetic control, - microfabricated membrane valves, or various other approaches to microfluidic transport and control. The combination of highly ordered flow and precise manipulation allows one to carry out diverse synthetic and analytical methods with remarkable control., Despite the near universal need for the preparation of standard samples, little work has been done to miniaturize and to expedite this process.
Approaches have included variously configured splitter channels - and differential metering of multiple inputs into addressable microfabricated assay wells. Each of these approaches to serial dilution requires N independent outputs (splitter branches, end reactors, etc.) for N consecutive dilutions, making them unsuitable for executing an arbitrary number of dilutions. The ideal circuit would automate sample and diluent metering and mixing, while scaling to an arbitrary number of serial dilutions.
A microfluidic mixing loop addresses mixing requirements by reducing effective diffusion lengths, while providing a compact geometry for manipulating nanoliter volumes. A microfluidic serial dilution circuit that implements these advantageous mixing and scaling characteristics and incorporates sample metering elements has been designed, fabricated, and characterized. It is compact and does not geometrically constrain the number of possible serial dilutions. Precise metering of the sample carryover fraction and rapid, reproducible mixing of the diluent with the carryover are achieved in the same structure.
The device is computer controlled, and the preparation of successive serial dilutions is fully automated. Because the circuit employs microfluidic pumping, serially diluted sample aliquots can easily be routed from the dilution circuit to other microfluidic components, such as a separation channel or microreactor. Microdevice Fabrication and Design The three-layer glass-PDMS-glass sandwich structure was fabricated as described previously., Features on the fluidic and manifold glass wafer layers were isotropically etched to a depth of 50 μm. The etched fluidic and manifold layers were visually aligned and reversibly bonded to one another with an intervening optically transparent PDMS membrane (250 μm thick, Rogers Corporation, Carol Stream, IL). Visual alignment and reversible bonding was performed in a laminar flow hood to minimize particulate contamination of the clean glass wafers and PDMS membrane.
Nylon tubing barbs (1/16″) were affixed to the fluidic chip surface at five pneumatic access holes to interface pneumatic control line tubing with the device. All reservoirs and vacuum access holes were drilled with 1.1-mm-diameter diamond-coated drill bits. A schematic of the microfluidic serial dilution circuit is shown in. Fluidic channels (black) are 300 μm wide, and valve deflection chambers (gray) are 1 mm in diameter. Both layers are 50 μm deep. All dimensions are after isotropic etching.
Two-way valves A, B, and C control fluid flow in the loop. Bus valves I and O control fluidic access to the input and output reservoirs, R i and R o, respectively. The loop remains continuous when the bus valves are closed, but fluid flow from R i and to R o is prevented.
The boxed inset to depicts the cross-sectional view of the glass-PDMS-glass sandwich structure. Schematic of the microfluidic serial dilution circuit. Fluidic channels are shown in black and pneumatic features are shown in gray. The input and output fluidic access reservoirs (1.1-mm diameter) are labeled R i and R o, respectively. The five membrane valve deflection chambers are labeled A, B, C, I, and O on their respective pneumatic lines. Valves A, B, and C are two-way valves and are continuous only when open.
Input and output valves I and O are bus valves, connecting R i and R o to the mixing loop. When open, I and O allow flow from R i and R o to and from the mixing loop. Fluidic continuity is preserved within the mixing loop even when I and O are closed. The boxed diagram depicts a cross section of the device at a two-way valve junction, showing the fluidic and manifold wafers, the PDMS membrane, the fluidic channel and discontinuity, and the corresponding valve displacement chamber. Pneumatic Control Computer-controlled pneumatic actuation of the membrane valves was accomplished using a TTL-driven vacuum solenoid valve array (HV010, Humphrey, Kalamazoo, MI).
On TTL low, the solenoid directs atmospheric pressure output, and the associated membrane valve rests in the closed state. On TTL high, the solenoid switches to vacuum and causes the associated membrane valve to deflect open. Download naruto shippuden 131 subtitle indonesia. The solenoid array is driven by the digital output of a NI6715 data acquisition PCMCIA card and PC laptop with software written in house (LabVIEW, National Instruments, Austin, TX). A sequence of valve states defines a pumping program.
A variable hold step interposed between states in the sequence is the valve actuation time. Three pumping programs were written to manipulate fluid in the serial dilution circuit. The valve sequences of each pumping program are written showing only the open valves at each step, and the hold step is indicated by a comma after each state in the sequence. For example, the program (AB, B) starts with valves A and B open and valves C, I, and O closed. This state is followed by a hold step, then valve A is closed leaving only B open.
The mix pumping program is the valve state sequence (A, AB, B, BC, C, AC). The flush pumping program is the valve state sequence (A, AB, B, BO, IO, IA). The prime pumping program is the valve state sequence (I, ACI, AC, ABCO, BO, O). Looping a pumping program results in continuous pumping., Each pumping program requires two input parameters for operation: the valve actuation time (in milliseconds) and the length of time the program is iterated (in seconds). Fluidic manipulation protocols are described in the text using the format: program(valve actuation time,iteration time), with valve actuation times given in milliseconds and iteration times given in seconds. For example, mix(80,60) indicates that the mix program is run with 80 ms valve actuation time, iterated for 60 s. Flow Visualization and Device Characterization Flow in the channels was visualized using a solution of fluorescein dye (10 μM in TAE) and a fiber-coupled epifluorescence microscope (488-nm laser excitation), which has been described.
Epifluorescence movies of the various pumping programs were acquired using a 12-bit CoolSnap FX CCD (10 fps, 50-ms exposure, 8 × 8 pixel binning, Roper Scientific, Tucson, AZ). The illumination area was ∼1.2 cm diameter and the power density was 1 mW/mm 2. Confocal fluorescence data were acquired using an inverted microscope fabricated in house. Laser excitation from a frequency-doubled diode laser was coupled into the optical detection train with a dichroic long-pass mirror (505DRLP, Omega Optical, Brattleboro, VT) and focused on the microfluidic channels with an infinite conjugate microscope objective (40× 0.6 NA, Newport, Irvine, CA).
Fluorescence was collected with the same objective, spectrally filtered with a bandpass filter (535DF60, Omega Optical), and focused with a 100-mm focal length achromatic lens on a 100- μm pinhole before impinging a photon counting PMT (H7827, Hamamatsu Corp., Japan). For all confocal fluorescence measurements, the detector was positioned in the fluidic channel region bounded by valves A and B.
Fluid handling characteristics of the device were quantitated using confocal fluorescence microscopy. The input reservoir, R i, was spotted with fluorescein solution and the circuit was run with prime(200,30) to prime with dye.
A syringe loaded with TAE buffer (the diluent) was used to rinse away residual dye solution in R i and to load diluent. This standard procedure was used to prepare the circuit for each of the following device characterization studies. The intrinsic carryover fraction (CF) for each serial dilution circuit was determined. The average fluorescence signal of the concentrated dye was measured, then the circuit was run with flush(100,60), and the average buffer background fluorescence signal was measured. Finally, the circuit was run with mix(100,120) to mix the carryover into the diluent. After mixing, the average fluorescence signal of the diluted dye was measured.
The ratio of the background-subtracted diluted dye signal to the dye concentrate signal is the CF. To demonstrate multiple serial dilutions of the same sample, a sample of 10 μM fluorescein was diluted in TAE using a mixing loop with CF of 0.2. To increase dynamic range, an OD 1 neutral density filter (Newport) was placed in line to measure the sample concentrate fluorescence intensity. Thereafter, the filter was removed and the fluorescence intensity of each consecutive dilution was measured as described above.
Fluidic handling reproducibility was evaluated by performing replicate dilutions. For each replicate, the circuit was prepared as described. Then the circuit was run with flush(100,90), followed by mix(100,120). Mixing was characterized by performing dilutions with variable valve actuation time during the mixing step. The circuit was primed as described, and mix(x,500) was initiated, where x was systematically varied from 300 ms to 50 ms. RESULTS AND DISCUSSION The serial dilution of an analyte can be automated and carried out on the nanoliter scale using an appropriately configured microfluidic mixing loop.
In-line computer-controlled membrane valves allow precise fluidic manipulation, automation, and parallelization. Fluidic operations, such as diluent flushing, mixing, and priming can be accurately and precisely performed without manual intervention, and performed simultaneously in many parallel circuits. A quantitative description of device performance was developed using epifluorescence flow visualization and confocal fluorescence microscopy. Epifluorescence visualization of the pumping programs flush and mix is presented in.
Diluent is pumped into the circuit through I, then through A and B, and finally out of the circuit through O. A plug of material in the region bounded by valves I and O and containing C is preserved by flush.
Frames 1 through 4 show TAE buffer (diluent) being pumping from R i to R o around the right side of the fluorescein dye-primed circuit. A plug of fluorescein dye (carryover) is preserved on the left (frame 4).
The carryover and diluent are mixed together in the mix operation by serially actuating valves A, B, and C while keeping valves I and O closed. Frames 5 through 8 show the carryover being mixed into the diluent as the fluid is cyclically pumped, and the fluorescence intensity in the loop homogenizes. Serial dilution circuit pumping program schematics and epifluorescence stills. Still frames are 50-ms exposures. The circuit is initially primed with fluorescein dye.
Fluid flow paths are indicated with gray arrows overlaid on the circuit schematic. The flush program is used for diluent flushing and carryover isolation, and is accomplished by serially actuating I, A, B, and O while keeping C closed. Buffer is pumped from R i to R o, clearing the right side of the mixing loop while isolating the carryover aliquot on the left side (frames 1−4). An example of an open valve can be seen in frame 2, in which B is open and the entire valve is filled with the concentrated dye solution. The mix program is used to mix the diluent and the isolated carryover by serially actuating A, B, and C while I and O are kept closed (frames 5−8). The output reservoir, R O, was manually evacuated in the time between frame 7 and frame 8 for the purpose of visualizing the fully mixed sample. Full movies are included in the.
A flush operation coupled to a mix operation constitutes a microfluidic serial dilution. Sample in the loop can be serially diluted many times to bring about consecutive serial dilutions of the concentrated sample. This concept is presented in. The detector was positioned between valves A and B (, inset) to observe three consecutive serial dilutions of fluorescein dye concentrate (300 nM). As the dye is cyclically pumped, the concentrated dye signal is acquired. Next, flush(100,60) and mix(100,120) are run sequentially to perform the serial dilution.
The measured fluorescence is reduced to background as the buffer diluent passes the detector during flush, then a mixing transient is observed during mix as the diluent and carryover mix. Once mixing is complete, the same program sequence is repeated to generate multiple serial dilutions (, inset). Quantitative evaluation of serial dilution. (A) Three consecutive serial dilutions of fluorescein dye solution (300 nM in TAE buffer) into TAE buffer were monitored using confocal fluorescence microscopy. The detector position is indicated in the inset circuit schematic.
The second and third dilutions are shown in the five-fold magnified inset. Serial dilutions were performed by executing flush(100,60) followed by mix(100,120). (B) A standard curve for 10 μM fluorescein was constructed from the average fluorescence intensity of the sample concentrate, and the intensity obtained after each of four consecutive six-fold dilutions.
Each data point represents the average of eight independent experiments. The log plot exhibits excellent linearity over the three detectable orders of magnitude (R 2 = 0.999). The construction of a complete series of standards based on a single 10 μM fluorescein standard solution is presented in. The log of the fluorescence intensity after each serial dilution was plotted as a function of the serial dilution cycle number, which is expected to be linear with slope proportional to the log of the carryover fraction (CF) of the circuit. The intrinsic CF for a circuit is determined by the fraction of the mixing loop bounded by valves I and O containing valve C. This fraction linearly depends on the angle θ subtended by the arc between valves I and O (, inset).
The CF of circuits with various θ was measured and plotted as a function of θ. Linear agreement of CF with θ is excellent (R 2 = 0.998). The error associated with each CF determination was 1.5%. During flush steps the carryover is still in contact with the flushing diluent stream, so carryover sample near the I and O valve boundaries may diffuse into the diluent stream. As the diluent flush time is increased, more sample diffuses out and the CF decreases. The dependence of CF on flush time was studied using fluorescein dye and buffer, and found to vary by 5% over the range of 30−300 s.
Dependence of carryover fraction on device geometry. The carryover fraction was related to the arc subtended by valves I and O. The inset indicates the angle measurement, θ.
CF = −0.02 + 0.005 θ; R 2 = 0.998. Microfluidic devices are characterized by the reproducibility of operations such as mixing and dilution because the flow regime is laminar. This concept is illustrated in. Replicate observations of a serial dilution conducted on two different devices demonstrate not only the reproducibility of dilutions performed in the same circuit, but also of dilutions performed on different devices. The inset of presents an overlay of the replicates.
Given identical fluidic programming, the rate of diluent flushing and the oscillations in the mixing transient are reproduced exactly between replicates. Mixing reproducibility. A solution of fluorescein dye was diluted using a circuit with a carryover fraction of 0.12. Two separate devices were operated with identical pumping parameters: flush(100,90), mix(100,120). The five profiles are offset by 200 CPS for clarity. The start of the flush and mix programs is indicated by arrows. The inset contains an overlay of the five replicates and a sample fit of an exponentially damped sinusoid.
Diluent flushing and mixing are highly reproducible, with mixing transients agreeing in fit within 1%. In order to study the reproducibility of the mixing transient quantitatively, a dampened sinusoid was fit to the data. The functional dependence of the damped sinusoid, Ae − kt sin( ω t) + b, contained least-squares fit parameters A, k, ω, and b, corresponding to the amplitude, damping factor, frequency, and offset after dilution, respectively. Typical R 2 values ranged from 0.90 to 0.98.
Parameter ω was fit with less than 0.3% least-squares error, and the frequency determined from fits of the five replicates agreed within 1%. A typical fit curve is shown in the inset, offset from the overlay.
Serial Dilution Lab Conclusion
Values of R 2 less than 0.95 are attributed to a relatively poor description of damping by the exponential term. Nonetheless, this procedure yielded excellent data on the transient frequency for the purpose of demonstrating the reproducibility of mixing. Sabki ba rate aayi doli tu bhi lana. The time required to mix the diluent plug into the carryover plug is influenced by the pumping rate, or valve actuation time, during cyclic mixing. Presents the dependence of the mixing transient morphology on the valve actuation time. As the valve actuation time is decreased from 300 ms to 50 ms, the linear flow velocity increases, and the mixing transient is compressed in time. As the two plugs are pumped through each other, mixing is expedited by the establishment of more diffusion planes.
The dependence of mixing time on valve actuation time can be determined qualitatively from. At 50 s, for example, the fluorescence intensity is still widely varying in the 300-ms case, while the signal has completely steadied in the 50-ms case. Mixing transients at variable valve actuation times. (A) Mixing transients were generated with variable actuation times and aligned to time t = 0. (B) Standard deviations as a function of time are plotted as solid lines, sampling valve actuation times of 300, 200, 100, and 50 ms. The standard deviation window width is the period of the oscillation for each transient.
A dashed line at σ win = 300 CPS indicates the threshold for complete mixing. Mixing times (.) measured at different valve actuation times are plotted discretely with respect to the left axis. Mixing times determined by this method exhibited ∼5% standard error. A quantitative study of mixing time is presented in. The standard deviation of an n-second-wide window, σ win, was plotted as a function of time to measure signal variance. The window width, n, was normalized by setting it equal to the transient period, 2π/ω, determined by fitting a damped sinusoid to each transient (described above).
Serial Dilution Microbiology
The deviation predictably drops as mixing proceeds. When the carryover and diluent are completely mixed, the standard deviation of the signal is limited by the shot noise of the detector, σ bkgd. The mixing time is the time required for σ win to reach 2σ bkgd. At this limit of detection, the observer is theoretically unable to differentiate between contributions to signal variance that arise systematically (as a result of incomplete mixing) versus those that arise randomly (as a result of shot noise). An analysis of mixing time as a function of valve actuation time, plotted discretely in, reveals that mixing is expedited as valve actuation time is decreased from 300 ms to 80 ms. The time required for complete mixing is minimized from 150 s to 22 s over the range of actuation times studied.
Further decreasing the valve actuation time from 80 ms to 50 ms did not significantly affect the mixing time. This agrees with measurements of linear flow rate as a function of valve actuation time; valve actuation appears to be limiting at valve actuation times shorter than 80 ms. The flow rate over the range of 80- to 50-ms valve actuation times gradually becomes independent of valve actuation time. Additionally, at higher flow velocities, transverse diffusion is limiting and the mixing time cannot be decreased absent a mechanism for establishing new boundary layers, for example by promoting torsional flow., Serial dilution is a common operation in chemical measurements. The construction of a series of standard samples can be time consuming and expensive, requiring many fluid metering steps and expending potentially valuable sample. The circuit described here carries out serial dilutions in 400 nL, though this is not a limit for circuit size.
In practice this circuit could be scaled down or up depending on the desired sample volume. Design constraints would include the valve dead volume and carryover channel volume. This microfluidic circuit can generate an entire standard curve with only the diluent as an input. The standards are prepared in nanoliter quantities, conserving reagent and allowing facile integration with on-chip analytical techniques. For example, on-chip capillary electrophoresis or liquid chromatography could be coupled to the output of this circuit, relying on integrated pumping for standard injection.
Importantly, this device can execute rapid and automated serial dilutions on the time scale of replication of a population of evolving RNA molecules, opening new avenues of inquiry in molecular evolution. CONCLUSIONS A microfluidic serial dilution circuit was developed that can perform multiple serial dilutions, with greatly increased speed and precision compared to manual pipeting.
Based on alterations of the circuit geometry, carryover fractions of 0.04−0.2 were demonstrated. The circuit requires only 400 nL of starting material to perform an arbitrary number of serial dilutions in a manner that can be integrated with other on-chip preparative or analytical steps. This circuit was developed to enable the automated serial dilution of a population of evolving RNA molecules, but is more generally applicable to almost any microfluidic architecture that involves serial dilution coupled to chemical synthesis or analysis.