This work was supported by Natural Sciences and Engineering Research Council of Canada (grant 402105), Canadian Foundation for Innovation (grant 129087) and Alberta Education and Training (grant AAETRCP-12-026-SEG) to RJC.

1. Calculations for Experimental Setup
<ol>
	<li>Calculate the volume of algal and/or bacterial cultures needed for controls that will be required for the entire experiment by using Equation 1: 
	<img alt="Equation 1" src="/files/ftp_upload/52455/52455equation1.1.jpg" /> 
	Where y equals the number of controls needed per day and z equals the number of days.</li>
	<li>Calculate the volume of algal and/or bacterial cultures that are needed for co-cultures for the experiment by using Equation 2: 
	<img alt="Equation 2" src="/files/ftp_upload/52455/52455equation1.2.jpg" /> 
	NOTE: It is possible to replace &#8216;co-culture&#8217; experiments with any compound screen; just adjust the final compound, solvent and algal concentrations to fit the experimental design.</li>
	<li>Use Equations 1 and 2 to calculate the final volume of early-exponential algal culture (commonly 5 days for ~104 cells/ml but this should be determined by performing an algal growth curve) required for the experiment (this volume is needed for step 2.3) using Equation 3: 
	<img alt="Equation 3" src="/files/ftp_upload/52455/52455equation1.3.jpg" /></li>
	<li>Use Equations 1 and 2 to calculate the final volume of 104 cfu/ml bacterial culture (or desired inoculation concentration) required for the experiment (this volume is needed for step 4.2) using Equation 4: 
	<img alt="Equation 4" src="/files/ftp_upload/52455/52455equation1.4.jpg" /> 
	NOTE: Use the additional 10 ml in steps 1.3 and 1.4 to account for pipetting error and any tests that are performed (e.g., WATER-PAM, flow cytometry, microscopy, etc.) on 0 d. Increase this volume if needed to fit the experimental design. 
	NOTE: For an example calculation of an 8-day experiment see Section 10. See supplemental table for media recipes.</li>
</ol>
2. Growing Algal Cells for Experimental Setup
<ol>
	<li>Isolate or obtain an actively growing axenic algal culture.</li>
	<li>Aseptically transfer 10% of the final volume of the algal culture into sterile algal medium (e.g., L1 or similar marine algal medium, see Materials and Equipment table), ensuring a 1:9 dilution of algal culture in fresh medium. Grow the diluted alga in a diurnal incubator using previously determined growth conditions for that strain (e.g., 18 &#176;C with a 16:8 hr light:dark cycle is commonly used).</li>
	<li>When the culture has reached early-exponential phase, re-culture the alga in the same sterile algal medium (e.g., L1 or similar marine algal medium), ensure the final concentration is a 1:9 dilution and that the final volume of algae is equal to the VA calculated (step 1.3). 
	NOTE: It is important to ensure these cultures are axenic. Test all algal media and algal stock bottles for contamination by plating a 20 &#956;l aliquot onto a general marine bacterial medium (e.g., Marine Broth 2216 supplemented with 1.5% agar or similar).</li>
	<li>Grow algal culture to early-exponential phase (~104 cells/ml). 
	NOTE: Every algal strain has a unique growth curve depending on culturing conditions. Early-exponential phase (~104 cells/ml) can be determined by performing an algal growth curve based on cell density (i.e. using flow cytometry or microscopy).</li>
</ol>
3. Preparing Bacterial Cells for Inoculation
<ol>
	<li>Pre-determine the bacterial concentration (cfu/ml) and optical density (OD) of the bacterium at stationary phase by doing a growth curve in the chosen bacterial medium and growth conditions (e.g., Marine Broth 2216 at 25 &#176;C and 160 rpm, or similar). Use this information to grow the cells (step 3.3) and later to dilute the cells correctly in step 3.7.</li>
	<li>Aseptically transfer an isolated and freshly grown bacterial colony from a 1.5% agar plate (e.g., Marine Broth 2216 supplemented with 1.5% agar or similar marine bacterial solid medium) into 5 ml of bacterial liquid medium (e.g., Marine Broth 2216 or similar marine bacterial medium).</li>
	<li>Grow the bacterium to stationary phase on a rolling drum or shaker (~12 - 36 hr, depending on the bacterium). Plan the experiment so that the bacterium reaches stationary phase (~108 - 109 cfu/ml) at the same time that the algal cells have reached the early-exponential phase (~104 cells/ml).</li>
	<li>Using a pipette, wash down any biofilm attached to the test tube into the medium. Pipette 1 ml of well-mixed bacterial culture into a sterile 1.5 ml microtube. Centrifuge for 1 min at 14,000 x g.</li>
	<li>Remove and dispose of the supernatant (bacterial media) without disrupting the pellet (bacterial cells). Add 1 ml of sterile algal media (e.g., L1 or similar) to the microtube (with pellet). Vortex microtube to resuspend pellet in algal media.</li>
	<li>Second wash: repeat step 3.5. 
	NOTE: It is critical to wash the bacterial cells with algal media in order to thoroughly remove all of the bacterial media, cell detritus, excreted proteins and small molecules from the cells prior to inoculating the algae with them as this could change the nutrient composition of the algal media or introduce bioactive molecules to the screen.</li>
	<li>Serially dilute the washed bacterial cells in algal media, to a final concentration that is 100-fold more concentrated than the desired final bacterial concentration (cfu/ml). Save the microtube containing cells that have been washed and diluted in algal media for step 4.2. 
	NOTE: Plan the experiment based on having a 1:1 ratio of algae to bacteria on 0 d. To do this the desired initial bacterial concentration for the experiment is 104 cfu/ml, so in step 3.7 the initial cells should be serially diluted to 106 cfu/ml.</li>
</ol>
4. Preparing Bacteria for Experimental Setup
<ol>
	<li>Prepare 4 sterile autoclaved glass conical flasks and label them: a) &#8216;algal control flask&#8217;, b) &#8216;diluted bacterial stock flask&#8217;, c) &#8216;bacterial control flask&#8217;, and d) &#8216;co-culture flask&#8217;.</li>
	<li>Dilute the bacterial suspension from step 3.7 1:99 with sterile algal media to a final volume = VB (step 1.4). Make the dilution in the flask labeled diluted bacterial stock (step 4.1). 
	NOTE: In the previous example (step 4.2), this 1:99 dilution gives a final concentration of 104 cfu/ml. Swirl flask to mix cells.</li>
	<li>Pipette Vcontrol (step 1.1) from the diluted bacterial stock and put it in the bacterial control flask.</li>
	<li>Pipette Vcontrol of sterile algal medium into the bacterial control flask (this is a 1:1 dilution). Swirl flask to mix cells and set aside for step 7.3.</li>
	<li>Pipette Vco-culture from the diluted bacterial stock flask to the co-culture flask, set flask aside for step 6.1. 
	NOTE: When doing multiple co-cultures on the same alga at once (i.e., a co-culture of two different bacterial isolates with one control) it is necessary to re-calculate the Vco-culture for each strain individually and then repeat step 4.5 for each co-culture separately. It is also necessary to increase the VA calculated in step 1.3 to include both co-cultures shown in Equation 5: 
	<img alt="Equation 5" src="/files/ftp_upload/52455/52455equation4.5.jpg" /></li>
</ol>
5. Preparing Algae for Experimental Setup
<ol>
	<li>Gently mix the early-exponential algal culture (from step 2.4) with a wide-mouth pipette tip until cells appear well mixed.</li>
	<li>Pipette Vcontrol from the algal stock bottle to the algal control flask. Gently pipette using a 10 ml pipette. Then return the algal stock bottle in the diurnal incubator.</li>
	<li>Pipette Vcontrol of sterile algal medium to the algal control flask (this is a 1:1 dilution). Swirl to mix flask and place in the diurnal incubator, until needed for step 7.4.</li>
</ol>
6. Preparing Experimental Co-culture
<ol>
	<li>Gently pipette Vco-culture from the algal stock flask into the bacterial co-culture flask from step 4.5. Return co-culture flask to the diurnal incubator until needed for step 7.5. 
	NOTE: The concentration of bacteria in the bacterial control should equal the bacterial concentration in the experimental co-culture. Similarly, the concentration of algae in the algal control should equal the concentration of algae in the experimental co-culture. By having the same initial bacterial and algal concentrations, the population density can be compared throughout the experiment.</li>
</ol>
7. Setting up Microtiter Plates
<ol>
	<li>Divide a sterile 48-well microtiter plate as per Figure 1. Label above outer wells containing either sterile diluent or non-photosynthetic samples. 
	NOTE: Randomization of the plate wells can be done for samples taken on the same day. For example, quadrant 1 will be sampled at 1 d in the experiment; the wells that should be randomized are B2 - 4 and C2 - 4. Leave the perimeter of the plate filled with sterile solution as shown in Figure 1.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/52455/52455fig1highres.jpg" /> 
Figure 1. Schematic representation of sample placement in a 48-well microtiter plate. Wells are to be filled as follows: columns 1 and 6, wells A through F (<img alt="Circle 1" src="/files/ftp_upload/52455/circle1.jpg" />) are filled with 1 ml 1x PBS (or other sterile solution/media). Rows A and F, wells 2 - 8 (<img alt="Circle 2" src="/files/ftp_upload/52455/circle2.jpg" />) are filled with 1 ml bacterial control; rows B and E wells 2 - 8 (<img alt="Circle 3" src="/files/ftp_upload/52455/circle3.jpg" />) are filled with 1 ml algal control; rows C and D, wells 2 - 8 (<img alt="Circle 4" src="/files/ftp_upload/52455/circle4.jpg" />) are filled with 1 ml co-culture. The plate is divided into 4 quadrants (A2, A5, D2, and D5) these quadrants are each specific sampling days 1 - 4, this should be randomized throughout plates and labeled accordingly. Within each day we advise randomizing the algal control (<img alt="Circle 5" src="/files/ftp_upload/52455/circle5.jpg" />) and co-culture (<img alt="Circle 6" src="/files/ftp_upload/52455/circle6.jpg" />) wells using a random number generator. Label the lid over the 1x PBS and/or bacterial control wells to prevent shading of algal cultures.
<ol start="2">
	<li>Pipette 1 ml 1x phosphate buffer solution (PBS, pH 7.4) or other sterile solution in the appropriate wells as indicated in Figure 1. Perform this slowly and with care. PBS will change the ionic strength of the algal and/or co-culture medium if it splatters in other wells so handle carefully.</li>
	<li>Pipette 1 ml of bacterial control culture into wells labeled bacterial control (Figure 1). Swirl the bacterial flask before pipetting each plate to avoid bacterial settling.</li>
	<li>Pipette 1 ml of algal control culture into wells labeled algal control using a wide mouth pipette tip (Figure 1). Swirl the algal flask as regularly as the bacterial flask. If the alga tends to sink or float, swirl as regularly as is needed to maintain a visually uniform culture.</li>
	<li>Using a wide mouth pipette tip, pipette 1 ml of co-culture in the appropriate wells (Figure 1). Swirling the co-culture flask as regularly as the algal flask.</li>
	<li>Seal each plate with parafilm and place in a diurnal incubator at the desired temperature and diurnal light cycle (18 &#176;C with a 16:8 hr light:dark cycle is commonly used). Leave the plates in the diurnal incubator until ready to take PAM readings (Section 8). Ensure all plates are oriented in the same direction to allow for consistent light exposure.</li>
	<li>Take a 20 &#956;l aliquot from the PBS, algal media, algal stock and algal control and drop plate onto an appropriate non-selective medium (e.g., Marine Broth 2216 supplemented with 1.5% agar) and incubate plates at 18 - 25 &#176;C for 72 hr to test for contamination. If growth appears in any of the solutions that should not contain bacteria, then these data should not be used.</li>
	<li>Take PAM fluorometry readings using the remaining sample from the algal control and co-culture flasks for the experimental 0 d PAM fluorometry reading (see step 8.1). Any other 0 d measurements should also be performed with the remaining algal control, bacterial control and co-culture samples.</li>
</ol>
8. Taking PAM Fluorometry Readings from Stock Samples
<ol>
	<li>Zero the WATER-PAM with sterile algal media in a clean cuvette before taking readings8.</li>
	<li>Pipette 300 &#181;l from algal control or co-culture flasks into a clean cuvette containing 2.7 ml of the same algal medium used in the experiment. Mix the sample and diluent gently with a wide mouth pipette tip.</li>
	<li>Wipe-off all fingerprints from outside of cuvette with a tissue before placing the cuvette in the WATER-PAM.</li>
	<li>Place cuvette into WATER-PAM. Cover the sample with the cap and allow it to dark-adapt for 3 min. Dark adaptation times vary depending on the species of algae and must be determined for the specific alga used in the experiment. Avoid lengthy dark adaptation times (&#62;20 min) by taking WATER-PAM readings during the middle of the dark cycle of the algal diurnal incubators16,17.</li>
	<li>After dark adaption, hit F0 button. If fluorescence readings are above 3,900, dilute sample 1:1 in algal medium. Dark-adapt for an additional 3 min and take a new reading. If the F0 or Fm readings are still above 3,900, continue to dilute the sample 1:1 in algal medium until the F0 and Fm readings are below 3,900. 
	NOTE: Make sure to account for these dilutions when recording final fluorescence: for instance if the algal sample is diluted 1:9 during the initial transfer from the well to the dilution tube, then the algal fluorescence reading of 500 should be multiplied by the inverse of the dilution factor (in this case 10) and the actual fluorescence of the tube is then 5,000.</li>
	<li>After setting F0, take a saturating pulse (SAT-Pulse) reading every 90 sec by hitting the SAT button to take Fm readings. The time interval between readings may be adjusted depending on algal strain. Discard sample.</li>
	<li>Repeat steps 8.1 - 8.6 for the remaining samples.</li>
</ol>
9. Taking PAM Fluorometry Readings from Microtiter Plates
<ol>
	<li>Label 6 sterile sample tubes of &#62;3 ml volume for wells B2 - 4 for the algal control and wells C2 - 4 for the bacterial co-culture (see plate layout in Figure 1).</li>
	<li>Aliquot 2.7 ml of sterile algal medium into each tube.</li>
	<li>Place tubes in diurnal incubator and allow them to acclimate to the temperature the alga was grown at for 30 min.</li>
	<li>Before removing the microtiter plate from the incubator ensure that light penetration into the dark acclimated wells is limited by covering the plate with aluminum foil (or similar), only remove the foil while actively transferring culture from the wells to the dilution tubes (steps 9.5 - 9.7).</li>
	<li>Aseptically mix the first microtiter plate well (well B2) with a wide mouth pipette tip by slowly pipetting up and down.</li>
	<li>Obtain the dilution tubes from the incubator and aseptically transfer 300 &#181;l from well B2 (Figure 1) to its corresponding sample tube with a wide mouth pipette tip (this is a 1:9 dilution of the sample in algal medium).</li>
	<li>Repeat steps 9.5 - 9.6 for the remaining wells (B3,4 and C2 - 4).</li>
	<li>Cover sample tubes with aluminum foil and return them to the diurnal incubator until ready for WATER-PAM.</li>
	<li>Perform WATER-PAM readings as described in steps 8.1 - 8.7. Seal the microtiter plate with parafilm before returning it to the incubator.</li>
	<li>Repeat readings at planned time intervals for the duration of the experiment. The frequency and length of sampling should be planned at the beginning of the experiment.</li>
</ol>
10. Sample Experiment
The sample experiment is a 10 day co-culture of a bacterium (Phaeobacter gallaeciensis BS107) and a microalga (Emiliania huxleyi strain (CCMP3266)). It includes an algal control, bacterial control, and a bacterial-algal experimental co-culture.
<ol>
	<li>Calculate volumes of bacterial and algal stocks required (steps 1.1 - 1.4) 
	<img alt="Equation 10" src="/files/ftp_upload/52455/52455equation10.1.jpg" /></li>
	<li>Prepare the algal stock, a VA of 40 ml, by inoculating 4 ml of alga in 36 ml medium and incubated until early-exponential growth is achieved with a cell density of ~104 cells/ml (steps 2.1 - 2.4).</li>
	<li>Prepare the bacterial stock flask by diluting 400 &#956;l of the 106 cfu/ml bacteria obtained in Section 3 to a VB of 40 ml with algal media (final concentration of bacteria will be 104 cfu/ml).</li>
	<li>Transfer half (20 ml) of the bacterial stock to the bacterial control flask and add 20 ml of algal media to make the bacterial control. Transfer half (20 ml) the algal stock to the algal control flask and add 20 ml of algal media to make up the algal control (Section 5). Transfer the remaining 20 ml of bacterial stock to the co-culture flask and mix with the remaining 20 ml of algal stock to establish the co-culture (Section 6).</li>
	<li>Pipette the 1x PBS (pH 7.4), controls, and co-culture in two pre-labeled microtiter plates (Figure 1), 1 ml per well. Use 3 ml of the remaining controls and co-cultures to make 0 d measurements (cfu counts, WATER-PAM (Section 8), microscopic observation of algal cell morphology, and algal cell fixation for flow cytometry).</li>
	<li>Take WATER-PAM readings for the algal control and co-culture once a day for 10 d, and always at the same time as the 0 d measurements were taken (Section 8).</li>
</ol>
11. Other Parameters of Interest
<ol>
	<li>Determine bacterial cfu concentration: perform a serial dilution of the bacterial control and co-culture in sterile 1x PBS (or similar), then drop plate onto Marine Broth 2216 supplemented with 1.5% agar (or similar) and observe the number of cfu that grow to determine the bacterial cfu/ml for each well18.</li>
	<li>To evaluate the algal cell concentration: fix algal control and co-culture with a 0.15% final concentration of glutaraldehyde. Incubate for 10 min in the dark, then flash freeze in liquid nitrogen, and store at -80 &#176;C. Process all samples on a flow cytometer (FACS Calibur or similar) to count algal cells.</li>
	<li>Observe algal cell morphology: algal cultures and bacterial-algal co-culture using microscopy (i.e., light microscopy, epifluorescence, or similar).</li>
</ol>

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Phycol. 35 (1), 69-74 (2000).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> WATER-PAM Chlorophyll Fluorometer. Instrument Description and Information for Users. , (2013).</li><li>Maxwell, K., Johnson, G. M., Heers, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chlorophyll+fluorescence--a+practical+guide.">Chlorophyll fluorescence--a practical guide.</a> J. Exp. Bot. 51 (345), 659-668 (2000).</li><li>Herigstad, B., Hamilton, M., Heersink, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=How+to+optimize+the+drop+plate+method+for+enumerating+bacteria.">How to optimize the drop plate method for enumerating bacteria.</a> J. Microbiol. 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Bot. 51 (345), 659-668 (2000).</li><li>Schreiber, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pulse-Amplitude-Modulation+(PAM)+Fluorometry+and+Saturation+Pulse+Method:+An+Overview.">Pulse-Amplitude-Modulation (PAM) Fluorometry and Saturation Pulse Method: An Overview.</a> Chlorophyll a Fluoresc. A Signat. Photosynth. , 279-319 (2004).</li><li>Roh&#225;&#269;ek, K., Bart&#225;k, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Technique+of+the+modulated+chlorophyll+fluorescence:+basic+concepts,+useful+parameters,+and+some+applications.">Technique of the modulated chlorophyll fluorescence: basic concepts, useful parameters, and some applications.</a> Photosynthetica. 37 (3), 339-363 (1999).</li><li>Da Silva, J. M., da Silva, A. B., P&#225;dua, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulated+chlorophyll+a+fluorescence:+a+tool+for+teaching+photosynthesis.">Modulated chlorophyll a fluorescence: a tool for teaching photosynthesis.</a> J. Biol. Educ. 41 (4), 178-183 (2007).</li><li>Vieira, S., Ribeiro, L., Jesus, B., Cartaxana, P., da Silva, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photosynthesis+assessment+in+microphytobenthos+using+conventional+and+imaging+pulse+amplitude+modulation+fluorometry.">Photosynthesis assessment in microphytobenthos using conventional and imaging pulse amplitude modulation fluorometry.</a> Photochem. Photobiol. 89 (1), 97-102 (2013).</li></ol>

The authors have nothing to disclose.

Algal growth in a miniaturized format.
The miniaturization of algal cultures to a 1 ml culture volume in a microtiter plate allows for the replication within an experiment to be increased. It is important to ensure the alga is healthy throughout an experiment; perform a growth curve (Figure 2), using the microtiter plate format to assess various algal media, to ensure the nutritional requirements of the alga are met. Additionally, it may be important to optimize the diurnal cycle...

Phytoplankton physiology has traditionally been studied in meso-scale experiments ranging from 20 ml in conical flasks to 5 L in carboys1&#8211;7. This experimental scale requires subsampling for experimental monitoring, as sacrificing replicate samples for each time point creates an unmanageable experimental setup.
The ability to increase the number of independent experiments while using the same diurnal incubator space by miniaturizing the experimental volume for algal physiology experiments will reduce or eliminate the limitations of subsampling and pseudo-replication from large volumes. A microtiter plate format has been developed for algal bioassays using a 1 ml culture volume for experimentally manipulating algae in variable conditions. This small experimental volume allows for the number of replicates to be increased, increases experimental reproducibility due to a decreased variability between replicate samples and experiments, and allows true replication while maintaining experimental controls (i.e., axenic algal cultures) for 140 days (Figure 2)12.
This microtiter plate format is easily adapted for a variety of experimental questions, such as: does a bacterium have a symbiotic, neutral or pathogenic interaction with its algal host? Is the addition of a compound stimulating or toxic to an alga? These and other questions can be addressed in a rapid high-throughput manner using this new format9&#8211;11.
A 48-well microtiter culture plate allows each 1 ml well to be an independent experimental setup that is sampled at a single time-point. Various parameters can be sampled from this 1 ml volume including, but not limited to: chlorophyll fluorescence and photochemical parameters using WATER-Pulse-Amplitude-Modulated (WATER-PAM) fluorometry (see Materials and Equipment table)13. WATER-PAM fluorometry is a rapid and non-invasive technique that can be used to monitor experiments performed with algae13. It allows measurement of photosynthetic efficiency and PSII health from a small culture volume (150 - 300 &#956;l of culture diluted in medium to a 2 - 4 ml volume for WATER-PAM)14,15. In addition to WATER-PAM fluorometry, this setup can be used to measure a variety of other parameters including, but not limited to: microscopy to visualize the bacteria attached to algal cells and changes in the algal cell morphology; bacterial colony forming unit (cfu) counts; and flow cytometry for algal cell counts and identifying subpopulations.

<table><tbody><tr><td>10 cu. ft. Diurnal incubator (6012-1)</td><td>Caron Corporate</td><td>112310-6012-1-11</td><td>&lt;a href=&quot;http://www.caronproducts.com&quot;&gt;www.caronproducts.com&lt;/a&gt;</td></tr><tr><td>Nunc EasYFlask 25 cm&lt;sup&gt;2&lt;/sup&gt;, Vent/Close Cap, 7 ml working volume, 200/cs</td><td>Thermo Fisher Scientific</td><td>N156340</td><td>&lt;a href=&quot;http://www.fishersci.ca&quot;&gt;www.fishersci.ca&lt;/a&gt;</td></tr><tr><td>Multiwell TC Plates &amp;ndash; 48-well</td><td>BD Biosciences Discovery Labware</td><td>353078</td><td>&lt;a href=&quot;http://www.bdbiosciences.com/&quot;&gt;www.bdbiosciences.com&lt;/a&gt;</td></tr><tr><td>P1000 Gilson The Pipetting Standard&amp;mdash;Gilson&#039;s Pipetman</td><td>Mandel Scientific Company Inc.</td><td>GF-F123602</td><td>&lt;a href=&quot;http://www.mandel.ca&quot;&gt;www.mandel.ca&lt;/a&gt;</td></tr><tr><td>P10mL Gilson The Pipetting Standard&amp;mdash;Gilson&#039;s Pipetman</td><td>Mandel Scientific Company Inc.</td><td>GF-F161201</td><td>&lt;a href=&quot;http://www.mandel.ca&quot;&gt;www.mandel.ca&lt;/a&gt;</td></tr><tr><td>Wide Orifice Tips nonsterile [100&amp;ndash;1,250 &amp;micro;l]</td><td>VWR International</td><td>89079-468</td><td>&lt;a href=&quot;http://www.ca.vwr.com&quot;&gt;www.ca.vwr.com&lt;/a&gt;</td></tr><tr><td>Ultrafine Tips nonsterile [100&amp;ndash;1,250 &amp;micro;l]</td><td>VWR International</td><td>89079-470</td><td>&lt;a href=&quot;http://www.ca.vwr.com&quot;&gt;www.ca.vwr.com&lt;/a&gt;</td></tr><tr><td>Finntip 10 ml [Vol: 1 - 10 ml]</td><td>Thermo Fisher Scientific</td><td>9402151</td><td>&lt;a href=&quot;http://www.fishersci.ca&quot;&gt;www.fishersci.ca&lt;/a&gt;</td></tr><tr><td>WATER-Pulse Amplitude Modulation (Water-ED)</td><td>Heinz Walz GmbH, Effeltrich, Germany</td><td>EDEE0232</td><td>&lt;a href=&quot;http://www.walz.com&quot;&gt;www.walz.com&lt;/a&gt;</td></tr><tr><td>15 mm diameter quartz glass cuvette (WATER-K)</td><td>Caron Corporate</td><td></td><td>&lt;a href=&quot;http://www.caronproducts.com&quot;&gt;www.caronproducts.com&lt;/a&gt;</td></tr><tr><td>Sodium chloride (crystalline/certified ACS), Fisher Chemical</td><td>Thermo Fisher Scientific</td><td>Thermo Fisher Scientific</td><td>&lt;a href=&quot;http://www.fishersci.ca&quot;&gt;www.fishersci.ca&lt;/a&gt;</td></tr><tr><td>BD Difco Marine Broth 2216</td><td>BD Biosciences Discovery Labware</td><td>BD Biosciences Discovery Labware</td><td>&lt;a href=&quot;http://www.bdbiosciences.com&quot;&gt;www.bdbiosciences.com&lt;/a&gt;</td></tr><tr><td>BD Bacto Agar</td><td>BD Biosciences Discovery Labware</td><td>BD Biosciences Discovery Labware</td><td>&lt;a href=&quot;http://www.bdbiosciences.com&quot;&gt;www.bdbiosciences.com&lt;/a&gt;</td></tr><tr><td>L1 Medium Kit, 50 L</td><td>NCMA [National Center for Marine Algae and Microbiota</td><td>NCMA [National Center for Marine Algae and Microbiota</td><td>&lt;a href=&quot;http://www.ncma.bigelow.org&quot;&gt;www.ncma.bigelow.org&lt;/a&gt;</td></tr></tbody></table>

a small volume bioassay to assess bacterial/phytoplankton co-culture using water-pulse-amplitude-modulated (water-pam) fluorometry

Conventional methods for experimental manipulation of microalgae have employed large volumes of culture (20 ml to 5 L), so that the culture can be subsampled throughout the experiment1&#8211;7. Subsampling of large volumes can be problematic for several reasons: 1) it causes variation in the total volume and the surface area:volume ratio of the culture during the experiment; 2) pseudo-replication (i.e., replicate samples from the same treatment flask8) is often employed rather than true replicates (i.e., sampling from replicate treatments); 3) the duration of the experiment is limited by the total volume; and 4) axenic cultures or the usual bacterial microbiota are difficult to maintain during long-term experiments as contamination commonly occurs during subsampling.
The use of microtiter plates enables 1 ml culture volumes to be used for each replicate, with up to 48 separate treatments within a 12.65 x 8.5 x 2.2 cm plate, thereby decreasing the experimental volume and allowing for extensive replication without subsampling any treatment. Additionally, this technique can be modified to fit a variety of experimental formats including: bacterial-algal co-cultures, algal physiology tests, and toxin screening9&#8211;11. Individual wells with an alga, bacterium and/or co-cultures can be sampled for numerous laboratory procedures including, but not limited to: WATER-Pulse-Amplitude-Modulated (WATER-PAM) fluorometry, microscopy, bacterial colony forming unit (cfu) counts and flow cytometry. The combination of the microtiter plate format and WATER-PAM fluorometry allows for multiple rapid measurements of photochemical yield and other photochemical parameters with low variability between samples, high reproducibility and avoids the many pitfalls of subsampling a carboy or conical flask over the course of an experiment.

WATER-PAM fluorometry readings.
WATER-Pulse-Amplitude-Modulated (PAM) fluorometry is a quick and efficient method to determine the fluorescence (a proxy for chlorophyll content) and photosynthetic yield (PSII health) of algal cultures. The PAM WinControl software generates a spreadsheet of raw data values for (the following are the basic parameters for dark adapted algal samples):
&#160;&#160;&#160;&#160;F0 = fluorescence of dark-adapted cells
<p class="...

Watch this Scientific Journal Video about A Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated WATER-PAM Fluorometry at JoVE.com

A Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated WATER-PAM Fluorometry

The goal of this procedure is to demonstrate the reproducibility and adaptability of using a microtiter plate format for microalgal screening. This rapid screen combines WATER-Pulse-Amplitude-Modulated (WATER-PAM) fluorometry to measure photosynthetic yield as an indicator of Photosystem II (PSII) health with small volume bacterial-algal co-cultures.

A Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated (WATER-PAM) Fluorometry

Conventional methods for experimental manipulation of microalgae have employed large volumes of culture (20 ml to 5 L), so that the culture can be ...

a-small-volume-bioassay-to-assess-bacterialphytoplankton-co-culture

Research

JoVE Journal

Environment

10.5K Views.  University of Alberta. The goal of this procedure is to demonstrate the reproducibility and adaptability of using a microtiter plate format for microalgal screening. This rapid screen combines WATER-Pulse-Amplitude-Modulated (WATER-PAM) fluorometry to measure photosynthetic yield as an indicator of Photosystem II (PSII) health with small volume bacterial-algal co-cultures.

Video: A Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated WATER-PAM Fluorometry

Co-culture Models of Pseudomonas aeruginosa Biofilms Grown on Live Human Airway Cells

Bacterial biofilms have been associated with a number of different human diseases, but biofilm development has generally been studied on non-living surfaces. In this paper, we describe protocols for forming Pseudomonas aeruginosa biofilms on human airway epithelial cells (CFBE cells) grown in culture. In the first method (termed the Static Co-culture Biofilm Model), P. aeruginosa is incubated with CFBE cells grown as confluent monolayers on standard tissue culture plates. Although the bacterium is quite toxic to epithelial cells, the addition of arginine delays the destruction of the monolayer long enough for biofilms to form on the CFBE cells. The second method (termed the Flow Cell Co-culture Biofilm Model), involves adaptation of a biofilm flow cell apparatus, which is often used in biofilm research, to accommodate a glass coverslip supporting a confluent monolayer of CFBE cells. This monolayer is inoculated with P. aeruginosa and a peristaltic pump then flows fresh medium across the cells. In both systems, bacterial biofilms form within 6-8 hours after inoculation. Visualization of the biofilm is enhanced by the use of P. aeruginosa strains constitutively expressing green fluorescent protein (GFP). The Static and Flow Cell Co-culture Biofilm assays are model systems for early P. aeruginosa infection of the Cystic Fibrosis (CF) lung, and these techniques allow different aspects of P. aeruginosa biofilm formation and virulence to be studied, including biofilm cytotoxicity, measurement of biofilm CFU, and staining and visualizing the biofilm.

Co-culture Models of Pseudomonas aeruginosa Biofilms Grown on Live Human Airway Cells

This paper describes different methods of growing Pseudomonas aeruginosa biofilms on cultured human airway epithelial cells. These protocols can be adapted to study different aspects of biofilm formation, including visualization of the biofilm, staining of the biofilm, measuring the colony forming units (CFU) of the biofilm, and studying biofilm cytotoxicity.

Bacterial biofilms have been associated with a number of different human diseases, but biofilm development has generally been studied on non-living ...

Immunology and Infection

Unraveling the Unseen Players in the Ocean - A Field Guide to Water Chemistry and Marine Microbiology

Here we introduce a series of thoroughly tested and well standardized research protocols adapted for use in remote marine environments. The sampling protocols include the assessment of resources available to the microbial community (dissolved organic carbon, particulate organic matter, inorganic nutrients), and a comprehensive description of the viral and bacterial communities (via direct viral and microbial counts, enumeration of autofluorescent microbes, and construction of viral and microbial metagenomes). We use a combination of methods, which represent a dispersed field of scientific disciplines comprising already established protocols and some of the most recent techniques developed. Especially metagenomic sequencing techniques used for viral and bacterial community characterization, have been established only in recent years, and are thus still subjected to constant improvement. This has led to a variety of sampling and sample processing procedures currently in use. The set of methods presented here provides an up to date approach to collect and process environmental samples. Parameters addressed with these protocols yield the minimum on information essential to characterize and understand the underlying mechanisms of viral and microbial community dynamics. It gives easy to follow guidelines to conduct comprehensive surveys and discusses critical steps and potential caveats pertinent to each technique.

Here, we present a comprehensive protocol to assess the organic and inorganic nutrient availability and the abundance and structure of microbial and viral communities in remote marine environments.

Here we introduce a series of thoroughly tested and well standardized research protocols adapted for use in remote marine environments. The sampling ...

Microtiter Dish Biofilm Formation Assay

Biofilms are communities of microbes attached to surfaces, which can be found in medical, industrial and natural settings. In fact, life in a biofilm probably represents the predominate mode of growth for microbes in most environments. Mature biofilms have a few distinct characteristics. Biofilm microbes are typically surrounded by an extracellular matrix that provides structure and protection to the community. Microbes growing in a biofilm also have a characteristic architecture generally comprised of macrocolonies (containing thousands of cells) surrounded by fluid-filled channels. Biofilm-grown microbes are also notorious for their resistance to a range of antimicrobial agents including clinically relevant antibiotics.
The microtiter dish assay is an important tool for the study of the early stages in biofilm formation, and has been applied primarily for the study of bacterial biofilms, although this assay has also been used to study fungal biofilm formation. Because this assay uses static, batch-growth conditions, it does not allow for the formation of the mature biofilms typically associated with flow cell systems. However, the assay has been effective at identifying many factors required for initiation of biofilm formation (i.e, flagella, pili, adhesins, enzymes involved in cyclic-di-GMP binding and metabolism) and well as genes involved in extracellular polysaccharide production. Furthermore, published work indicates that biofilms grown in microtiter dishes do develop some properties of mature biofilms, such a antibiotic tolerance and resistance to immune system effectors.
This simple microtiter dish assay allows for the formation of a biofilm on the wall and/or bottom of a microtiter dish. The high throughput nature of the assay makes it useful for genetic screens, as well as testing biofilm formation by multiple strains under various growth conditions. Variants of this assay have been used to assess early biofilm formation for a wide variety of microbes, including but not limited to, pseudomonads, Vibrio cholerae, Escherichia coli, staphylocci, enterococci, mycobacteria and fungi.
In the protocol described here, we will focus on the use of this assay to study biofilm formation by the model organism Pseudomonas aeruginosa. In this assay, the extent of biofilm formation is measured using the dye crystal violet (CV). However, a number of other colorimetric and metabolic stains have been reported for the quantification of biofilm formation using the microtiter plate assay. The ease, low cost and flexibility of the microtiter plate assay has made it a critical tool for the study of biofilms.

The assay describes a rapid means to measure early biofilm formation in bacteria and fungi. This method uses a microtiter plate as the substratum for microbial biofilm formation, and the biofilm is visualized using crystal violet strain. The assay provides either a qualitative or quantitative assay for early biofilm formation.

Biofilms are communities of microbes attached to surfaces, which can be found in medical, industrial and natural settings.  In fact, life in a biofilm ...

Fluorescence detection methods for microfluidic droplet platforms

The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that accompany system miniaturisation. Advantages include the ability to efficiently process nano- to femoto- liter volumes of sample, facile integration of functional components, an intrinsic predisposition towards large-scale multiplexing, enhanced analytical throughput, improved control and reduced instrumental footprints.1
In recent years much interest has focussed on the development of droplet-based (or segmented flow) microfluidic systems and their potential as platforms in high-throughput experimentation.2-4 Here water-in-oil emulsions are made to spontaneously form in microfluidic channels as a result of capillary instabilities between the two immiscible phases. Importantly, microdroplets of precisely defined volumes and compositions can be generated at frequencies of several kHz. Furthermore, by encapsulating reagents of interest within isolated compartments separated by a continuous immiscible phase, both sample cross-talk and dispersion (diffusion- and Taylor-based) can be eliminated, which leads to minimal cross-contamination and the ability to time analytical processes with great accuracy. Additionally, since there is no contact between the contents of the droplets and the channel walls (which are wetted by the continuous phase) absorption and loss of reagents on the channel walls is prevented.
Once droplets of this kind have been generated and processed, it is necessary to extract the required analytical information. In this respect the detection method of choice should be rapid, provide high-sensitivity and low limits of detection, be applicable to a range of molecular species, be non-destructive and be able to be integrated with microfluidic devices in a facile manner. To address this need we have developed a suite of experimental tools and protocols that enable the extraction of large amounts of photophysical information from small-volume environments, and are applicable to the analysis of a wide range of physical, chemical and biological parameters. Herein two examples of these methods are presented and applied to the detection of single cells and the mapping of mixing processes inside picoliter-volume droplets. We report the entire experimental process including microfluidic chip fabrication, the optical setup and the process of droplet generation and detection.

Droplet-based microfluidic platforms are promising candidates for high throughput experimentation since they are able to generate picoliter, self-compartmentalized vessels inexpensively at kHz rates. Through integration with fast, sensitive and high resolution fluorescence spectroscopic methods, the large amounts of information generated within these systems can be efficiently extracted, harnessed and utilized.

The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that ...

Bioengineering

Characterization of Aquatic Biofilms with Flow Cytometry

Biofilms are dynamic consortia of microorganism that play a key role in freshwater ecosystems. By changing their community structure, biofilms respond quickly to environmental changes and can be thus used as indicators of water quality. Currently, biofilm assessment is mostly based on integrative and functional endpoints, such as photosynthetic or respiratory activity, which do not provide information on the biofilm community structure. Flow cytometry and computational visualization offer an alternative, sensitive, and easy-to-use method for assessment of the community composition, particularly of the photoautotrophic part of freshwater biofilms. It requires only basic sample preparation, after which the entire sample is run through the flow cytometer. The single-cell optical and fluorescent information is used for computational visualization and biological interpretation. Its main advantages over other methods are the speed of analysis and the high-information-content nature. Flow cytometry provides information on several cellular and biofilm traits in a single measurement: particle size, density, pigment content, abiotic content in the biofilm, and coarse taxonomic information. However, it does not provide information on biofilm composition on the species level. We see high potential in the use of the method for environmental monitoring of aquatic ecosystems and as an initial biofilm evaluation step that informs downstream detailed investigations by complementary and more detailed methods.

Flow cytometry in combination with visual clustering offers an easy-to-use and fast method for studying aquatic biofilms. It can be used for biofilm characterization, detection of changes in biofilm community structure, and detection of abiotic particles embedded in the biofilm.

Biofilms are dynamic consortia of microorganism that play a key role in freshwater ecosystems. By changing their community structure, biofilms respond ...

Fluorescently Labeled Bacteria as a Tracer to Reveal Novel Pathways of Organic Carbon Flow in Aquatic Ecosystems

Elucidating trophic interactions, such as predation and its effects, is a frequent task for many researchers in ecology. The study of microbial communities has many limitations, and determining a predator, prey, and predatory rates is often difficult. Presented here is an optimized method based on the addition of fluorescently labelled prey as a tracer, which allows for reliable quantitation of the grazing rates in aquatic predatory eukaryotes and estimation of nutrient transfer to higher trophic levels.

Presented here is a protocol for a single-cell, epifluorescence microscopy-based technique to quantify grazing rates in aquatic predatory eukaryotes with high precision and taxonomic resolution.

Elucidating trophic interactions, such as predation and its effects, is a frequent task for many researchers in ecology. The study of microbial ...

A Platform of Anti-biofilm Assays Suited to the Exploration of Natural Compound Libraries

Biofilms are regarded as one of the most challenging topics of modern biomedicine, and they are potentially responsible for over 80% of antibiotic-tolerant infections. Biofilms have displayed an exceptionally high tolerance for chemotherapy, which is thought to be multifactorial. For instance, the matrix provides a physical barrier that decreases the penetration of antibiotics into the biofilm. Also, cells within the biofilms are phenotypically diverse. Likely, biofilm resilience arises from a combination of these and other, yet unknown, mechanisms. All of the currently existing antibiotics have been developed against single-cells (planktonic) bacteria. Therefore, so far, a very limited repertoire of molecules exists that can selectively act on mature biofilms. This situation has driven a progressive paradigm shift in drug discovery, in which searching for anti-biofilms has been urged to occupy a more prominent place. An additional challenge is that there are a very limited number of standardized methods for biofilm research, especially those that can be used for large-throughput screening of chemical libraries. Here, an experimental anti-biofilm platform for chemical screening is presented. It uses three assays to measure biofilm viability (with resazurin staining), total biomass (with crystal violet staining), and biofilm matrix (using a wheat germ agglutinin, WGA-fluorescence-based staining of the poly-N-acetyl-glucosamine, PNAG, fraction). All the assays were developed using Staphylococcus aureus as the model bacteria. Examples of how the platform can be used for primary screening as well as for functional characterization of identified anti-biofilm hits are presented. This experimental sequence further allows for the classification of the hits based upon the measured end-points. It also provides information on their mode of action, especially on long-term versus short-term chemotherapeutic effects. Thus, it is very advantageous for the quick identification of high-quality hit compounds that can serve as starting points for various biomedical applications.

Biofilm infections show high tolerance towards chemotherapy. No single assay captures the complexity of biofilms. Instead, complementary assays are needed. We present a screening platform (developed for S. aureus) that combines assays for viability, biomass, and biofilm matrix. It allows anti-biofilm drug discovery, including the assessment of long-term chemotherapeutic effects.

Biofilms are regarded as one of the most challenging topics of modern biomedicine, and they are potentially responsible for over 80% of ...

Operation of Laboratory Photobioreactors with Online Growth Measurements and Customizable Light Regimes

The laboratory study of microalgae can be experimentally challenging. In addition to the cultivation requirements of non-photosynthetic microorganisms, phototrophs also require illumination. Routinely, researchers seek to provide custom light supplies, i.e., vary the light intensity and time over which it is delivered. Such flexibility is difficult with standard benchtop lights. Usually, cultivation studies also require growth comparisons between experimental treatments. Frequently, growth is assessed over an extended duration, e.g., multiple times a day over a week-long trial. Manual measurements can be time-consuming and lack data resolution. Therefore, photobioreactors (PBRs) with automatic growth monitoring and customizable light supply are useful for replicated experiments with multiple treatments. The current work presents the design, construction, and operation of laboratory PBRs. The materials are easily sourced and relatively inexpensive. The design could be duplicated with moderate skill. Each structure has a footprint of ~40 cm2 and hosts three 1 L glass bottles for triplicate replication. Bottles rest upon platforms containing magnetic stirrers and are arranged vertically within a 1 m high and 15 cm diameter polyvinyl chloride (PVC) pipe. The pipe interior is lined with light-emitting diodes (LEDs). These LEDs produce continuous light intensities from 0&#8211;2400 &#181;mol photons m-2 s-1 of photosynthetically active radiation (PAR). Users design a custom lighting program. The light intensity can be adjusted each second or held constant for longer durations. Oxygen produced from photosynthesis exits each bottle via a one-way volumetric gas sensor. Software is used to record gas sensor data. The amount of oxygen produced can be correlated to biomass growth. If biomass samples are required, a syringe can be used to extract culture. The method is suited for microalgae grown with bicarbonate as the carbon source. These PBRs are valuable to a laboratory that requires replicated experiments, light regime flexibility, and continuous high-resolution growth data.

This publication describes the design of laboratory photobioreactors (PBRs) with customizable light regimes. The growth of cyanobacteria or microalgae, using bicarbonate as their carbon source, is monitored continuously by measuring volumetric oxygen production. These PBRs facilitate rapid, replicated laboratory growth comparisons with little user intervention during experiments.

The laboratory study of microalgae can be experimentally challenging. In addition to the cultivation requirements of non-photosynthetic microorganisms, ...

Biology

Measuring Photophysiology of Attached Stage of Colacium sp. by a Cuvette-Type Fast Repetition Rate Fluorometer

Fast repetition rate fluorometer (FRRf) is a beneficial method for measuring photosystem II (PSII) photophysiology and primary productivity. Although FRRf can measure PSII absorption cross-section (&#963;PSII), maximum photochemical efficiency (Fv/Fm), effective photochemical efficiency (Fq&#8242;/Fm&#8242;), and non-photochemical quenching (NPQNSV) for various eukaryotic algae and cyanobacteria, almost all FRRf studies to date have focused on phytoplankton. Here, the protocol describes how to measure PSII photophysiology of an epizoic alga Colacium sp.&#160;Ehrenberg 1834 (Euglenophyta), in its attached stage (attached to zooplankton), using cuvette-type FRRf. First, we estimated the effects of substrate zooplankton (Scapholeberis mucronata O.F. M&#252;ller 1776, Cladocera, Daphniidae) on baseline fluorescence and &#963;PSII, Fv/Fm, Fq&#8242;/Fm&#8242;, and NPQNSV of planktonic Colacium sp. To validate this methodology, we recorded photophysiology measurements of attached Colacium sp. on S. mucronata and compared these results with its planktonic stage. Representative results showed how the protocol could determine the effects of calcium (Ca) and manganese (Mn) on Colacium sp. photophysiology and identify the various effects of Mn enrichment between attached and planktonic stages. Finally, we discuss the adaptability of this protocol to other periphytic algae.

Measuring Photophysiology of Attached Stage of Colacium sp. by a Cuvette-Type Fast Repetition Rate Fluorometer

Fast repetition rate fluorometer (FRRf) is a beneficial method for measuring photosystem II photophysiology and primary productivity. Here we describe a protocol to measure PSII photophysiology of epizoic alga, Colacium sp. on substrate zooplankton using cuvette-type FRRf.

Fast repetition rate fluorometer (FRRf) is a beneficial method for measuring photosystem II (PSII) photophysiology and primary productivity. Although FRRf ...

A Peg Plate Biofilm Assay for Screening Antibacterial Agents

Take a fresh 96-well plate, and add 100 microliters of CRPMI equilibrated to room temperature to each well of rows B to H. Dilute up to four test compounds separately in 1.0 milliliter of CRPMI to 2x the desired final concentration. Add 200 microliters of compound one in wells A1 to A3, 200 microliters of compound two in wells A4 to A6, 200 microliters of compound three in wells A7 to A9, and 200 microliters of compound four in wells A10 to A12.
Serially dilute the test compounds using a multichannel pipette. Begin by transferring 100 microliters from row A to row B, and mix. In that manner, continue to transfer 100 microliters well to well. Mix after each transfer and stop in row F. After mixing, expel 100 microliters from row F into a waste container. Do not transfer any liquid to rows G and H. Finally, add 100 microliters of CRPMI to each well of the plate using a multichannel pipette, to bring the final volume to 200 microliters.
Add 200 microliters of phosphate-buffered saline to a fresh sterile 96-well plate that features wells with dimensions identical to those of the biofilm initiation plate. Next, remove the plastic bag and paraffin film from the incubated biofilm initiation plate. Lift the lid straight up, avoiding contact between the pegs and the plate bottom as this may damage the biofilm. Keep the bottom part for subsequent OD600 analysis.
Rinse off planktonic cells by placing the peg lid onto the plate containing PBS. Submerge pegs for 5 seconds, lift them out of PBS, and submerge once more, being careful not to hit the pegs against the well sides. Place the rinsed peg lid into the challenge plate. Avoid unnecessary contact between pegs and the bottom plate, as this will damage the biofilm leading to inconsistent results.
Seal the plate with paraffin film, and place into a sealable plastic bag as before. Incubate the plate without shaking for 24 hours at 37 degrees Celsius in a 5% carbon dioxide incubator. Take the bottom of the biofilm initiation plate and resuspend the bacteria in each well using a multichannel pipette. Measure the OD600 with a suitable microplate reader to confirm growth occurring in all wells, but the sterility controls in row H.
Use a plate reader equipped with an incubation chamber, and capable of taking kinetic fluorescence readings for detecting resazurin conversion. Set up a 24-hour kinetic fluorescence measurement using a 530-nanometer excitation and 590-nanometer emission. Set the chamber temperature to 37 degrees Celsius and have the plate read every 20 minutes.
Set the plate reader to measure from the bottom of the plate, according to the manufacturer&#39;s instructions. Prepare to wash the plate by adding 200 microliters of PBS to each well of a sterile 96-well plate with a multichannel pipette. Then, prepare biofilm recovery medium by adding 400 microliters of 0.8 milligram per milliliter resazurin stock solution to 20 milliliters of CRPMI medium, to a final concentration of 16 micrograms per milliliter resazurin.
Next, add 150 microliters of biofilm recovery medium with a multichannel pipette, to each well of a fresh 96-well plate. Transfer the challenge plate from the incubator into a biosafety cabinet, and carefully remove the plastic bag and sealing film. Remove the peg lid from the challenge plate and rinse off planktonic cells in PBS as before, keeping the bottom of the challenge plate for subsequent OD600 analysis.
After rinsing, place the peg lid into the plate bottom containing the biofilm recovery medium. Wrap the side of the plate with paraffin film, being careful not to obstruct the bottom of the perimeter wells. Immediately after wrapping the plate, place it on the plate reader, and start a kinetic read over a 24-hour period by initiating the previously made resazurin kinetic protocol.

A Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated (WATER-PAM) Fluorometry

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