Name: fluorescently labeled bacteria as a tracer to reveal novel pathways of organic carbon flow in aquatic ecosystems
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This study was supported by the Czech Science Foundation under the research grant 13-00243S and 19-16554S awarded to K. &#352;. and D. S., respectively. This article was also supported by the project &#34;Biomanipulation as a tool for improving water quality of dam reservoirs&#34;&#160;(No CZ.02.1.01/0.0/0.0/16_025/0007417), funded by the European Regional Development Fund, in Operational Programme Research, Development and Education.

1. Sample collection
<ol>
	<li>Collection of reservoir water sample: the first case study (Exp I; lower natural in situ predator and prey abundance system)
	<ol>
		<li>Collect water samples from the desired location at a suitable depth. Keep the samples in a temperature-controlled cooler filled at in situ temperature (avoiding temperature shock; it should be noted that uptake rates of protists are temperature-dependent) during transport to the laboratory. 
		NOTE: Our sampling was conducted in the meso-eutrophic canyon-shaped &#344;&#237;mov reservoir (South Bohemia, volume 34.5 x 106 m3, maximal depth 43 m, mean retention time 100 days, dimictic). The sampling site was located at a 30 m depth, close to the dam. A mixed sample at a 0.5 m depth was collected by a 2-l Friedinger sampler.</li>
		<li>Continue to section 1.2 as soon as possible.</li>
	</ol>
	</li>
	<li>Collection of fluid from the traps of carnivorous Utricularia reflexa plants: the second case study (Exp II; a system with high predator and prey abundances, extremely small sample volume)
	<ol>
		<li>Gently shake the plants underwater, remove from the cultivation container, and place them on absorbent material to absorb excess water. Depending on the robustness of the plants and size of the traps, choose 8-10 plant individuals.</li>
		<li>Divide each shoot into roughly equal parts by counting leaf nodes bearing trapping organs. Each shoot segment will serve to collect mixed samples representing young, middle-aged, and old traps.</li>
		<li>Attach a thin glass capillary and Eppendorf vial for sample collection to a peristaltic pump. Proceed to insert the capillary tip into the trap opening. Using the vacuum pump, suck out all the fluid from each trap until 900 &#177; 100 &#181;L of trap fluid is collected for each trap age category.</li>
		<li>Use triplicate subsamples of 200 &#181;L of the pooled trap fluid for protistan grazing experiments. Process these immediately as detailed in section 3. Preserve the remaining &#177;300 &#181;L of the sample for all other analyses of microbial components living in the fluid, as detailed below (section 2).</li>
		<li>Proceed immediately to section 2.</li>
	</ol>
	</li>
</ol>
2. Fixation of collected samples
<ol>
	<li>Exp I and II: fix water/trap fluid sub-samples for bacterial enumeration (section 4; approximately 20 mL and 0.3 mL, respectively) with formaldehyde for at least 1 h to obtain a 2% final volume:volume concentration in each sample. 
	NOTE: Handle formaldehyde exclusively in the fume hood, and wear gloves at all times while manipulating samples.</li>
</ol>
3. Sample filtration
<ol>
	<li>Sample dilution (Exp I): no dilution is needed for the reservoir water samples. (Exp II): dilute the trap fluid sample 10x-100x with particle-free MQ water to achieve a suitable distribution of the target microbes on filter surfaces prior to counting via epifluorescence microscopy.</li>
	<li>Filter 1-2 mL (Exp I) of the reservoir water or 10-30 &#956;L (Exp II) of the subsample of the trap fluid for bacterial counting onto black 0.2 &#181;m pore-size filters, using a filtration funnel (25 mm diameter).</li>
	<li>Stain the filters with DAPI (4&#39;,6-diamidino-2&#39;-phenylindole dihydrochloride, 0.2% final concentration) for 4 min. 
	NOTE: Avoid skin and working surface contamination, and wear gloves.</li>
	<li>Place the filter with concentrated microbes onto a drop of immersion oil (for fluorescent microscopy) on a microscope slide. Place another oil drop onto the center of the filter and cover with a coverslip, making sure that the oil is distributed evenly.</li>
	<li>At this point, preferably process samples immediately, or alternatively, store them in the freezer (-20 &#176;C) for several weeks to months until further analysis.</li>
</ol>
4. Enumeration of bacterial numbers on the filters
<ol>
	<li>Place the slide under the epifluorescence microscope (with the filter set corresponding to the fluorochrome DAPI). Place a 10 x 10 counting grid into one of the oculars. Move the slide to a random position.</li>
	<li>Quantify bacterial cells (blue fluorescence) in the area of the counting grid (under the magnification 1000x). In the counts, include the cells crossing the left and upper edges of the counting grid, while excluding those located across the right and lower edges.</li>
	<li>Move to another random position and repeat enumeration on at least 10-15 counting grids, amounting to 500 cells counted in total.</li>
	<li>Establish a conversion factor for a given microscope and magnification, based on knowledge of the ratio of the area of one grid to the total effective filtration area of the filter. Then, divide the total number of cells counted by the number of grids counted, yielding an average number of bacteria per grid.</li>
	<li>Multiply the latter parameter by the conversion factor established and normalize the resulting number per mL of a sample (depending on the volume of sample filtered) to obtain the total bacterial abundance per mL.</li>
</ol>
5. Determining protistan abundance
<ol>
	<li>Fix the water sample (Exp I) or trap fluid (Exp II) subsamples with either glutaraldehyde (1% final concentration, more suitable for samples with the presence of chlorophyll containing particles to be processed days to weeks after fixation) or using the formol-thiosulfate decolorization technique specified below. 
	NOTE: Both preservation techniques prevent egestion of the ingested material from food vacuoles of protists8.</li>
	<li>Regarding the formol-thiosulfate decolorization technique, add 100 &#181;L/1 &#181;L of Lugol&#8217;s solution into 20 mL/200 &#181;L of the water/trap fluid subsample (Exp I/Exp II, respectively).</li>
	<li>Follow immediately with the addition of 0.5 mL/50 &#181;L of borate-buffered formalin then 20 &#181;L/2 &#181;L of 3% sodium thiosulphate (Exp I/Exp II, respectively). 
	NOTE: The sodium thiosulphate decolorizes the yellow color of Lugol to enable observation of cells under the epifluorescence microscope8.</li>
	<li>Filter a known volume of sample (depending on the number of target protists) onto 1 &#181;m pore-size black polycarbonate filters.</li>
	<li>Estimate the number of the protist species is by counting at least 200 cells under magnification 600x using the counting grid (see above).</li>
	<li>Reduce the volume of the sample under a vacuum in the filtration funnel by low vacuum to approximately 2 mL. Then, release the under-pressure and add the DAPI fluorochrome (4&#39;,6-diamidino-2-phenylindole dihydrochloride13, 0.2% final concentration) for 2 min.</li>
</ol>
6. Determining community structure of ciliates in plankton samples
NOTE: Ciliate communities in freshwater habitats are highly diverse14,15,16,18, and their microscopic determination is challenging. Sorting the ciliate groups into functional guilds10,14,16,17 allows for more detailed analysis of different ciliate groups as pelagic bacterivores.
<ol>
	<li>Evaluate ciliate community structure by combining the following:
	<ol>
		<li>DAPI-stained samples in epifluorescence microscopy (to localize the ciliate cells with bright fluorescence of macro- and micro-nuclei of different sizes and morphology) combined with the uptake of fluorescently labeled bacteria (FLB8; for details, see below), tracking the ability of ciliates to feed on bacteria.</li>
		<li>Live sample observation in selected cases17,19. For more details of the above approaches and criteria used for grouping ciliates into different taxonomic categories, see previous publications16,17. 
		NOTE: Cited studies have indicated that among ciliates, omnivorous species from Stichotrichia(genera&#160;Halteria and Pelagohalteria) and Oligotrichia (namely Rimostrombidium spp.) are the most important pelagic consumers of bacterioplankton in a vast majority of freshwater habitats10,17,18.</li>
	</ol>
	</li>
</ol>
7. Estimating ciliate grazing rates
<ol>
	<li>Calculate the uptake rates of ciliates on bacteria based on changes in the average number of tracer [i.e., FLB8 per ciliate related to time of incubation (5-15 min)] and tracer amount of FLB added, accounting at most&#160;for 5%-15% of total bacteria.</li>
	<li>To compare uptake rates among different protistan species, normalize the uptake rates as the number of bacteria per ciliate per hour, with calculations based on the actual incubation time and proportion of tracer FLB added. 
	NOTE: The general scheme of the FLB method application to estimate both cell- or species-specific and bulk bacterivory rates in natural samples is depicted in Figure 1.</li>
	<li>Preparation of FLB from bacterial strains indigenous to a freshwater environment8
	<ol>
		<li>Select a suitable size (mean cell volume = MCV) and morphology of bacteria so that they effectively mimic the typical sizes of bacterioplankton/bacterial cells in the aquatic system being investigated. 
		NOTE: For Exp I, a mixture of isolated strains from the study location of the genus Limnohabitans and Polynucleobacter was used (i.e., typical, highly abundant bacterioplankton in lakes and ponds)20. For details on morphology and sizes of the strains, see previous publications3,17,18,21.</li>
		<li>Harvest bacterial cells from culture by centrifugation (5,000 x g) for 15 min at the early stationary phase and mix them at a numerical ratio that yields MCV &#177; SD of the cells in the mixture corresponding to the typical MCV of bacteria at the chosen location.</li>
		<li>Suspend the pellets in 10 mL of phosphate-buffered saline (PBS; pH = 9).</li>
		<li>Add 2 mg of the yellow-green fluorescing dye 5-(4,6-dichlorotriazin-2-yl) aminofluorescein (DTAF, binds to proteins) to the cell suspension in the phosphate-saline buffer, and incubate in a 60 &#176;C water bath for 2 h.</li>
		<li>After the incubation, centrifuge the cells down, decant the DTAF solution, and wash and centrifuge 3x with PBS.</li>
		<li>After the final wash, re-suspend the cells in 20 mL of the PPi-saline buffer.</li>
		<li>Vortex the FLB suspension and pipette 1.5 mL aliquots into 2 mL cryo-vials, then keep frozen (at -20 &#176;C) in PPi-saline buffer until use.</li>
		<li>Pre-filter PPi-saline buffer through a 0.2 &#181;m polycarbonate filter for use in the next step.</li>
		<li>To determine FLB concentration, transfer a small aliquot (usually 20-40 &#181;L) to 2 mL of particle-free PPi-saline buffer, sonicate at 30 W for several 2 s bursts, and filter onto 0.2 &#181;m polycarbonate black filter for enumeration via epifluorescence microscopy (1,000X magnification) under optical filter settings for DTAF (448 nm/520-540 nm).</li>
	</ol>
	</li>
	<li>Tracer technique for estimation of ciliate bacterivory
	<ol>
		<li>For grazing experiments in natural planktonic habitats, dispense 300 mL samples into well-rinsed 1 L flasks and incubate at in situ temperature for 15 min (to allow protists to recover from the handling shock).</li>
		<li>Add FLB tracers to constitute 5%-15% of total bacteria, with the amounts added depending upon the season and water temperature. 
		NOTE: There is a very broad, season-dependent spectrum in ciliate species-specific uptake rates spanning several orders of magnitude (i.e., from 101-104 bacteria ciliate-1 per hour)10,16,17,18,22,23,24.</li>
		<li>In periods of enhanced occurrence of ciliates with high uptake rates (usually during the summer), also run a parallel incubation with very low FLB additions, constituting only 2%-4% of total bacteria to avoid excessive loading of ciliate vacuoles by tracer FLB (see examples in Figure 2).</li>
		<li>Incubate ciliate/plankton samples with FLB for 5-15 min.</li>
		<li>There are two possibilities for sample fixation to prevent egestion of the ingested material from food vacuoles of protists8. Terminate the incubations by the addition of 1% glutaraldehyde (final concentration that is more suitable for samples with chlorophyll-containing particles, such as algae). Alternatively, use 100 &#181;L/10 &#181;L of Lugol&#39;s solution into 20 mL/200 &#181;L of water/trap fluid subsample, followed immediately by the addition of 0.5 mL/10 &#181;L of borate-buffered formalin then 200 &#181;L/2 &#181;L of 3% sodium thiosulphate (Exp I/Exp II, respectively).</li>
	</ol>
	</li>
	<li>After adding the fixative, let the samples rest for at least 1 h in the dark at 4 &#176;C to ensure thorough preservation of ciliate cells.</li>
	<li>Take natural plankton subsamples from 4-30 mL/10-30 &#956;L (Exp I/Exp II, respectively; the volume depends on ciliate abundance) and stain with DAPI (final concentration 0.2% wt/vol; for details, see step 3.2 above).</li>
	<li>Pass through 1 &#181;m black filters and inspect via epifluorescence microscopy to count ciliates (600x magnification) and enumerate the number of FLB tracers ingested (mostly at 1000x magnification) as detailed in previous publications2,17. Inspect samples within 7 days after preservation.</li>
	<li>To estimate total protozoan/species-specific grazing, multiply average uptake rates of all ciliates, or of only the ciliate species as detected by in situ abundance.</li>
	<li>Example of the calculation of per-cell uptake rates from in situ data from the &#344;&#237;mov water reservoir is described as follows:
	<ol>
		<li>Assume that the bacterial concentration is 3.55 x 106 bacteria/mL and tracer FLB added is 0.25 x 106 FLB/mL, which yields a sum of 3.8 x 106 bacteria/mL of total bacterial particles (natural bacteria + FLB = 100% of prey particles) available to phagotrophic protists in the natural sample. 
		NOTE: The FLB tracers added thus represent 6.58% (a project of 0.25/0.038) of total bacterial particles. The average number of FLB per Halteria sp. is 6.2 FLB in 5 min incubations.</li>
		<li>To normalize uptake per hour, use the following calculation: (6.2 x 12)/(6.58/100) = 1131 bacteria per Halteria cell/h. 
		NOTE: For more examples of distributions of individual uptake rates of Halteria sp. detected under variable water temperatures, amounts (as percents) of tracer FLB added, and different incubation times with FLB, see Figure 3.</li>
	</ol>
	</li>
</ol>

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T., Kratina, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diet+tracing+in+ecology:+Method+comparison+and+selection.">Diet tracing in ecology: Method comparison and selection.</a> Methods in Ecology and Evaluation. 9, 278-291 (2018).</li><li>Beisner, B. E., Grossart, H. P., Gasol, 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=A+guide+to+methods+for+estimating+phago-mixotrophy+in+nanophytoplankton.">A guide to methods for estimating phago-mixotrophy in nanophytoplankton.</a> Journal of Plankton Research. , 1-13 (2019).</li><li>Dolan, J. D., &#352;imek, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Processing+of+ingested+matter+in+Strombidium+sulcatum,+a+marine+ciliate+(Oligotrichida).">Processing of ingested matter in Strombidium sulcatum, a marine ciliate (Oligotrichida).</a> Limnology and Oceanography. 42, 393-397 (1997).</li><li>Massana, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Grazing+rates+and+functional+diversity+of+uncultured+heterotrophic+flagellates.">Grazing rates and functional diversity of uncultured heterotrophic flagellates.</a> The ISME Journal. 3, 588-596 (2009).</li><li>Gruj&#269;i&#263;, V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryptophyta+as+major+freshwater+bacterivores+in+experiments+with+manipulated+bacterial+prey.">Cryptophyta as major freshwater bacterivores in experiments with manipulated bacterial prey.</a> The ISME Journal. 12, 1668-1681 (2018).</li></ol>

The authors have nothing to disclose.

Deciphering trophic interaction in aquatic systems is always challenging28, especially at the nano-plankton scales involving protists and their prey, bacteria. When it comes to nutrient uptake pathways and quantification, the application of methods successfully used at higher trophic levels is less possible, due to the high complexity of biotic interactions. These include, for example, stable isotope labeling approaches. This protocol shows the advantages of using epifluorescence microscopy and fl...

Heterotrophic prokaryotes are a key biological component in aquatic systems and account for a significant fraction of plankton biomass1,2,3. Factors that control their abundance, diversity, and activity are crucial for understanding their role in biogeochemical cycling (i.e., the fate of organic carbon and other nutrients and flow of energy from prokaryotes to higher trophic levels). Protozoan grazing is one of these important factors. Bacterivory&#160;of heterotrophic nanoflagellates and ciliates imposes a strong top-down control over prokaryotic abundance, community function, structure, diversity, and even cellular morphology and growth rate of particular bacterial groups4,5,6. In some systems, protists serve as the major cause of bacterial mortality6,7.
The standard approach used to assess protozoan bacterivory, which has been used for some time now, involves the use of fluorescently labeled bacteria (FLB) as prey analogues and epifluorescence microscopy. Cell-specific uptake rates can be determined by quantifying the number of labeled prey particles in protistan food vacuoles over a selected time course8. There are several advantages to this approach. Tracer is added to natural samples with natural predator and prey assemblages. There is minimum sample manipulation prior to incubation, minimum sample alteration by the added FLB tracer, and incubation times are short to ensure sound results obtained under close to in situ conditions. Alternatively, in environments with low numbers of bacterivorous protists or zooplankton (e.g., offshore marine systems), disappearance rates of FLB added to samples in low amounts (2%-3% tracer) can be detected via flow cytometry in long-term (12-24 h) incubation experiments. Then, numbers of FLB at the start and end points (integrating the impact of all bacterivores) are quantified by flow cytometry (for details, see previous publication9). However, such a parameter only represents total aggregated bacterivory rates that cannot be directly attributed to any particular protistan and zooplankton grazer groups or species.
Overall, quantifying the protistan species- or morphotype-specific bacterial mortality rates in the aquatic environment accurately and with ecological meaning can be challenging. Some protists are selective grazers, and the size and cell shape of the added FLB tracer may distort natural rates of prey ingestion10,11. Moreover, protistan activity and metabolism are highly temperature-sensitive12; therefore, the amount of added FLB tracer needs to be carefully manipulated for each individual sample type (not only based on the natural abundance, size, and morphology of bacteria and prevailing types of bacterivores, but also on temperature). Most studies focus on bulk protistan grazing activity; however, the bacterivory of specific protistan species often holds a much higher information value and may be preferable. In this case, taxonomic knowledge of the protist species present in a sample and understanding of their behavior is needed. Hence, considerable amounts of time and labor are required to obtain sound results on species-specific rates of bacterivory attributable to a particular protistan group or species.
Despite these difficulties, this approach remains the most suitable tool currently available to assess protistan bacterivory in natural settings. Presented here is a comprehensive, easy-to-follow method for using FLB as a tracer in aquatic microbial ecology studies. All of the mentioned problematic aspects of the approach are accounted for and an improved workflow is described, with two experiments from contrasting environments as well as contrasting ciliate species as examples.
The first case study was conducted in an epilimnetic environment from the mesotrophic &#344;&#237;mov water reservoir in the Czech Republic, which shows grazer and bacterial abundances comparable to most surface freshwater bodies (cf.5,7). The second case study was conducted in the highly specific environment inside traps of the aquatic carnivorous plant Utricularia reflexa, which hosts extremely high numbers of both grazing mixotrophic ciliates (Tetrahymena utriculariae) and bacterial cells. Calculations of cell-specific grazing rates and bacterial standing stocks in both sample types are shown. A range of ecological interpretations of the results is then discussed, and examples of possible follow-up studies are finally suggested.

<table><tbody><tr><td>0.2-&amp;micro;m pore-size filters&amp;nbsp;</td><td>SPI supplies, https://www.2spi.com/</td><td>B0225-MB</td><td>Black, polycarbonate track etch membrane filters, diameter approprite for filtering apparatus used</td></tr><tr><td>5-(4,6-dichlorotriazin-2-yl) aminofluorescein (DTAF)</td><td>Any brand</td><td></td><td></td></tr><tr><td>Automatic pipettes with adjustable volumes&amp;nbsp;</td><td>Any brand, various sizes</td><td></td><td></td></tr><tr><td>Centrifuge</td><td>22 000 x g</td><td></td><td></td></tr><tr><td>Cryovials</td><td>Any brand, 2 mL size</td><td></td><td></td></tr><tr><td>DAPI (4&amp;acute;,6-Diamidino-2&amp;acute;-phenylindole dihydrochloride)</td><td>Any brand&amp;nbsp;</td><td></td><td>1 mg ml&lt;sup&gt;-1&lt;/sup&gt;</td></tr><tr><td>Epiflorescence microscope</td><td>Magnification from 400 x up to 1000 x</td><td></td><td></td></tr><tr><td></td><td>Filters appropriate for viewing in the DAPI and DTAF range</td><td></td><td></td></tr><tr><td></td><td>Counting grid in one of the oculars</td><td></td><td></td></tr><tr><td>Filtering apparatus</td><td>Usually with a diameter of 25 mm&amp;nbsp;</td><td></td><td></td></tr><tr><td>Formaldehyde</td><td>A brand for microscopy</td><td></td><td></td></tr><tr><td>Glutaraldehyde</td><td>A brand for microscopy</td><td></td><td></td></tr><tr><td>Immersion oil for microscopy</td><td>Specific oil with low fluorescence</td><td></td><td></td></tr><tr><td>Lugol&amp;acute;s solution</td><td>Any brand or see comment</td><td></td><td>Make an alkaline Lugol&#039; solution as follows: Solution 1 - dissolve&amp;nbsp; 10 g of potassium iodide in 20 ml in MQ water, then add 5 g of iodine. Solution 2 - add 5 g of sodium acetate&amp;nbsp; to 50 ml of MQ water. Add the solution 2 to the solution 1 and thoroughly mix</td></tr><tr><td>Methanol stabilized formalin</td><td>Any brand available for microscopy purposes</td><td></td><td></td></tr><tr><td>Microscope slides and cover slips</td><td>Any brand produced for microscopy purposes&amp;nbsp;</td><td></td><td></td></tr><tr><td>MQ water for diluting samples</td><td>Any brand&lt;br/&gt; &amp;nbsp;</td><td></td><td></td></tr><tr><td>Phosphate-buffered saline (PBS; pH = 9)</td><td>Any brand</td><td></td><td>0.05 M Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;-NaCl solution, adjusted to pH 9</td></tr><tr><td>PP&lt;sub&gt;i&lt;/sub&gt;-saline buffer</td><td>Any brand</td><td></td><td>0.02 M Na&lt;sub&gt;4&lt;/sub&gt;P&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;7&lt;/sub&gt;-NaCl solution. Add 0.53 g Na&lt;sub&gt;4&lt;/sub&gt;P&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;7&lt;/sub&gt; to 100 ml of MQ water plus 0.85 g NaCl&amp;nbsp;</td></tr><tr><td>Sampling device&amp;nbsp;</td><td>Appropriate for obtaining representative sample&amp;nbsp;</td><td></td><td>e.g. Friedinger sampler for lake plankton</td></tr><tr><td>Sodium thiosulfate solution</td><td>Any brand</td><td></td><td>3% solution is used in the protocol</td></tr><tr><td>Sonicator</td><td>Any brand</td><td></td><td>30 W</td></tr><tr><td>Vortex</td><td>Any brand allowing&amp;nbsp; thorough mixing of the solutes and samples</td><td></td><td></td></tr><tr><td>Water bath</td><td>Any brand allowing temperature to be maintained at 60 &amp;deg;C</td><td></td><td></td></tr></tbody></table>

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.

Example experiment I was run in &#344;&#237;mov water reservoir (South Bohemia, CZ), which is a natural site with lower natural in situ predator and prey abundance. Representative data is reported for the omnivorous ciliate species Halteria grandinella, which is an abundant and efficient grazer of picoplankton (&#60;2 &#181;m) particles10,16,17,18,<sup class...

Watch this Scientific Journal Video about Fluorescently Labeled Bacteria as a Tracer to Reveal Novel Pathways of Organic Carbon Flow in Aquatic Ecosystems at JoVE.com

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

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 ...

fluorescently-labeled-bacteria-as-tracer-to-reveal-novel-pathways

Universidad de Cartagena UDEC

Research

JoVE Journal

Environment

7.0K Views.  Biology Centre CAS. 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.

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

Monitoring Spatial Segregation in Surface Colonizing Microbial Populations

Microbes provide an intriguing system to study social interaction among individuals within a population. The short generation times and relatively simple genetic modification procedures of microbes facilitate the development of the sociomicrobiology field. To assess the fitness of certain microbial species, selected strains or their genetically modified derivatives within one population, can be fluorescently labelled and tracked using microscopy adapted with appropriate fluorescence filters. Expanding colonies of diverse microbial species on agar media can be used to monitor the spatial distribution of cells producing distinctive fluorescent proteins.
Here, we present a detailed protocol for the use of green- and red-fluorescent protein producing bacterial strains to follow spatial arrangement during surface colonization, including flagellum-driven community movement (swarming), exopolysaccharide- and hydrophobin-dependent growth mediated spreading (sliding), and complex colony biofilm formation. Non-domesticated isolates of the Gram-positive bacterium, Bacillus subtilis can be utilized to scrutinize certain surface spreading traits and their effect on two-dimensional distribution on the agar-solidified medium. By altering the number of cells used to initiate colony biofilms, the assortment levels can be varied on a continuous scale. Time-lapse fluorescent microscopy can be used to witness the interaction between different phenotypes and genotypes at a certain assortment level and to determine the relative success of either.

The role of assortment (spatial segregation) in evolutionary scenarios can be examined using simple microbial systems in the laboratory that allow controlled adjustment of spatial distribution. By modifying the founder cell density, various assortment levels can be visualized using fluorescently labelled bacterial strains in the colony biofilms of Bacillus subtilis.

Microbes provide an intriguing system to study social interaction among individuals within a population. The short generation times and relatively simple ...

Genetics

A Whole Cell Bioreporter Approach to Assess Transport and Bioavailability of Organic Contaminants in Water Unsaturated Systems

Bioavailability of contaminants is a prerequisite for their effective biodegradation in soil. The average bulk concentration of a contaminant, however, is not an appropriate measure for its availability; bioavailability rather depends on the dynamic interplay of potential mass transfer (flux) of a compound to a microbial cell and the capacity of the latter to degrade the compound. In water-unsaturated parts of the soil, mycelia have been shown to overcome bioavailability limitations by actively transporting and mobilizing organic compounds over the range of centimeters. Whereas the extent of mycelia-based transport can be quantified easily by chemical means, verification of the contaminant-bioavailability to bacterial cells requires a biological method. Addressing this constraint, we chose the PAH fluorene (FLU) as a model compound and developed a water unsaturated model microcosm linking a spatially separated FLU point source and the FLU degrading bioreporter bacterium Burkholderia sartisoli RP037-mChe by a mycelial network of Pythium ultimum. Since the bioreporter expresses eGFP in response of the PAH flux to the cell, bacterial FLU exposure and degradation could be monitored directly in the microcosms via confocal laser scanning microscopy (CLSM). CLSM and image analyses revealed a significant increase of the eGFP expression in the presence of P. ultimum compared to controls without mycelia or FLU thus indicating FLU bioavailability to bacteria after mycelia-mediated transport. CLSM results were supported by chemical analyses in identical microcosms. The developed microcosm proved suitable to investigate contaminant bioavailability and to concomitantly visualize the involved bacteria-mycelial interactions.

A whole cell bioreporter assay with Burkholderia sartisoli RP037-mChe was developed to detect fractions of an organic contaminant (i.e., fluorene) available for bacterial degradation after active transport by mycelia bridging air-filled pores in a water unsaturated model system.

Bioavailability of contaminants is a prerequisite for their effective biodegradation in soil. The average bulk concentration of a contaminant, however, is ...

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of sexually transmitted infection and also the primary cause of preventable blindness in the world. An essential determinant for successful infection of host cells by Chlamydia is the bacterium's ability to manipulate host cell signaling from within a novel, vacuolar compartment called the inclusion. From within the inclusion, Chlamydia acquire nutrients required for their 2-3 day developmental growth, and they additionally secrete a panel of effector proteins onto the cytosolic face of the vacuole membrane and into the host cytosol. Gaps in our understanding of Chlamydia biology, however, present significant challenges for visualizing and analyzing this intracellular compartment. Recently, a reverse-imaging strategy for visualizing the inclusion using GFP expressing host cells was described. This approach rationally exploits the intrinsic impermeability of the inclusion membrane to large molecules such as GFP. In this work, we describe how GFP- or mCherry-expressing host cells are generated for subsequent visualization of chlamydial inclusions. Furthermore, this method is shown to effectively substitute for costly antibody-based enumeration methods, can be used in tandem with other fluorescent labels, such as GFP-expressing Chlamydia, and can be exploited to derive key quantitative data about inclusion membrane growth from a range of Chlamydia species and strains.

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

A live cell fluorescent protein based method for illuminating cellular vacuoles (inclusions) containing Chlamydia is described. This strategy enables rapid, automated determination of Chlamydia infectivity in samples and can be used to quantitatively investigate inclusion growth dynamics.

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of ...

Immunology and Infection

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular level. To date the shear complexity of biological systems has caused precise single-molecule experimentation to be far too demanding, instead focusing on studies of single systems using relatively crude bulk ensemble-average measurements. However, many important processes occur in the living cell at the level of just one or a few molecules; ensemble measurements generally mask the stochastic and heterogeneous nature of these events. Here, using advanced optical microscopy and analytical image analysis tools we demonstrate how to monitor proteins within a single living bacterial cell to a precision of single molecules and how we can observe dynamics within molecular complexes in functioning biological machines. The techniques are directly relevant physiologically. They are minimally-perturbative and non-invasive to the biological sample under study and are fully attuned for investigations in living material, features not readily available to other single-molecule approaches of biophysics. In addition, the biological specimens studied all produce fluorescently-tagged protein at levels which are almost identical to the unmodified cell strains ("genomic encoding"), as opposed to the more common but less ideal approach for generating significantly more protein than would occur naturally ('plasmid expression'). Thus, the actual biological samples which will be investigated are significantly closer to the natural organisms, and therefore the observations more relevant to real physiological processes.

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Here we demonstrate the protocols for performing single-molecule fluorescence microscopy on living bacterial cells to enable functional molecular complexes to be detected, tracked and quantified.

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular ...

Biology

Bromodeoxyuridine (BrdU) Labeling and Subsequent Fluorescence Activated Cell Sorting for Culture-independent Identification of Dissolved Organic Carbon-degrading Bacterioplankton

Microbes are major agents mediating the degradation of numerous dissolved organic carbon (DOC) substrates in aquatic environments. However, identification of bacterial taxa that transform specific pools of DOC in nature poses a technical challenge. 
Here we describe an approach that couples bromodeoxyuridine (BrdU) incorporation, fluorescence activated cell sorting (FACS), and 16S rRNA gene-based molecular analysis that allows culture-independent identification of bacterioplankton capable of degrading a specific DOC compound in aquatic environments. Triplicate bacterioplankton microcosms are set up to receive both BrdU and a model DOC compound (DOC amendments), or only BrdU (no-addition control). BrdU substitutes the positions of thymidine in newly synthesized bacterial DNA and BrdU-labeled DNA can be readily immunodetected 1,2. Through a 24-hr incubation, bacterioplankton that are able to use the added DOC compound are expected to be selectively activated, and therefore have higher levels of BrdU incorporation (HI cells) than non-responsive cells in the DOC amendments and cells in no-addition controls (low BrdU incorporation cells, LI cells). After fluorescence immunodetection, HI cells are distinguished and physically separated from the LI cells by fluorescence activated cell sorting (FACS) 3. Sorted DOC-responsive cells (HI cells) are extracted for DNA and taxonomically identified through subsequent 16S rRNA gene-based analyses including PCR, clone library construction and sequencing.

Bromodeoxyuridine BrdU Labeling and Subsequent Fluorescence Activated Cell Sorting for Culture-independent Identification of Dissolved Organic Carbon-degrading Bacterioplankton

Environmental bacterioplankton are incubated with a model dissolved organic carbon (DOC) compound and a DNA labeling reagent, bromodeoxyuridine (BrdU). Afterward, DOC-degrading cells are separated from the bulk community based on their elevated BrdU incorporation using fluorescence activated cell sorting (FACS). These cells are then identified by subsequent molecular analyses.

Microbes are major agents mediating the degradation of numerous dissolved organic carbon (DOC) substrates in aquatic environments. However, identification ...

Kinetic Visualization of Single-Cell Interspecies Bacterial Interactions

Polymicrobial communities are ubiquitous in nature, yet studying their interactions at the single-cell level is difficult. Thus, a microscopy-based method has been developed for observing interspecies interactions between two bacterial pathogens. The use of this method to interrogate interactions between a motile Gram-negative pathogen, Pseudomonas aeruginosa and a non-motile Gram-positive pathogen, Staphylococcus aureus is demonstrated here. This protocol consists of co-inoculating each species between a coverslip and an agarose pad, which maintains the cells in a single plane and allows for visualization of bacterial behaviors in both space and time.
Furthermore, the time-lapse microscopy demonstrated here is ideal for visualizing the early interactions that take place between two or more bacterial species, including changes in bacterial species motility in monoculture and in coculture with other species. Due to the nature of the limited sample space in the microscopy setup, this protocol is less applicable for studying later interactions between bacterial species once cell populations are too high. However, there are several different applications of the protocol which include the use of staining for imaging live and dead bacterial cells, quantification of gene or protein expression through fluorescent reporters, and tracking bacterial cell movement in both single species and multispecies experiments.

This live-bacterial cell imaging protocol allows for visualization of interactions between multiple bacterial species at the single-cell level over time. Time-lapse imaging allows for observation of each bacterial species in monoculture or coculture to interrogate interspecies interactions in multispecies bacterial communities, including individual cell motility and viability.

Polymicrobial communities are ubiquitous in nature, yet studying their interactions at the single-cell level is difficult. Thus, a microscopy-based method ...

Whole&#45;Organ Pathogen Tracking via Fluorescent Reporter Quantification

High&#45;Resolution Three&#45;Dimensional Whole&#45;Organ Tomography of Microbial Infections

Most infections take place within three-dimensional host tissues with intricate anatomy and locally varying host physiology. The positioning of pathogen cells within this diverse environment significantly affects their stress levels, responses, fate, and contribution to the overall progression of the disease and treatment failure. However, due to the technical difficulties in locating &#181;m-sized pathogen cells within cm-sized host organs, this area of research has been relatively unexplored. Here, we present a method for addressing this challenge. We employ serial two-photon tomography and AI-enhanced image analysis to locate individual Salmonella cells throughout the entire spleen, liver lobes, and whole lymph nodes of infected mice. Using fluorescent reporters and in vivo antibody administration, the replication rate of single Salmonella cells, their local interaction with specific immune cells, and bacterial responses to antibiotics can be determined. These methodologies open avenues for a comprehensive examination of infections, their prevention, and treatment within the three-dimensional tissue context.

Author Spotlight: Unraveling Bacterial Responses to Antibiotics and Immune System in Tissues

Here, we describe a procedure that enables organ-wide detection of pathogenic bacteria during infection and quantification of fluorescent reporter activities.

Most infections take place within three-dimensional host tissues with intricate anatomy and locally varying host physiology. The positioning of pathogen ...

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

Live Cell Fluorescence Microscopy to Observe Essential Processes During Microbial Cell Growth

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of proteins which are essential for bacterial survival. A series of target-specific dyes can be used as probes to better understand these processes. Staining with lipophilic dyes enables the observation of membrane structure, visualization of lipid microdomains, and detection of membrane blebs. Use of fluorescent-d-amino acids (FDAAs) to probe the sites of peptidoglycan biosynthesis can indicate potential defects in cell wall biogenesis or cell growth patterning. Finally, nucleic acid stains can indicate possible defects in DNA replication or chromosome segregation. Cyanine DNA stains label living cells and are suitable for time-lapse microscopy enabling real-time observations of nucleoid morphology during cell growth. Protocols for cell labeling can be applied to protein depletion mutants to identify defects in membrane structure, cell wall biogenesis, or chromosome segregation. Furthermore, time-lapse microscopy can be used to monitor morphological changes as an essential protein is removed and can provide additional insights into protein function. For example, the depletion of essential cell division proteins results in filamentation or branching, whereas the depletion of cell growth proteins may cause cells to become shorter or rounder. Here, protocols for cell growth, target-specific labeling, and time-lapse microscopy are provided for the bacterial plant pathogen Agrobacterium tumefaciens. Together, target-specific dyes and time-lapse microscopy enable characterization of essential processes in A. tumefaciens. Finally, the protocols provided can be readily modified to probe essential processes in other bacteria.

Understanding the function of essential processes in bacteria is challenging. Fluorescence microscopy with target-specific dyes can provide key insights into microbial cell growth and cell cycle progression. Here, Agrobacterium tumefaciens is used as a model bacterium to highlight methods for live cell imaging for characterization of essential processes.

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of ...

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

A method is described for labeling individual messenger RNA (mRNA) transcripts in fixed bacteria for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH allows the measurement of cell-to-cell variability in mRNA copy number of genes of interest, as well as the subcellular location of the transcripts. The main steps involved are fixation of the bacterial cell culture, permeabilization of cell membranes, and hybridization of the target transcripts with sets of commercially available short fluorescently-labeled oligonucleotide probes. smFISH can allow the imaging of the transcripts of multiple genes in the same cell, with limitations imposed by the spectral overlap between different fluorescent markers. Following completion of the protocol illustrated below, cells can be readily imaged using a microscope coupled with a camera suitable for low-intensity fluorescence. These images, together with cell contours obtained from segmentation of phase contrast frames, or from cell membrane staining, allow the calculation of the mRNA copy number distribution of a sample of cells using open-source or custom-written software. The labeling method described here can also be applied to image transcripts with stochastic optical reconstruction microscopy (STORM).

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

This manuscript describes a method for labeling individual messenger RNA (mRNA) transcripts with fluorescently-labeled DNA probes, for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH is a visualization method that allows the simultaneous detection, localization, and quantification of single mRNA molecules in fixed individual cells.

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

Summary

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Chapters in this video

Monitoring Spatial Segregation in Surface Colonizing Microbial Populations

A Whole Cell Bioreporter Approach to Assess Transport and Bioavailability of Organic Contaminants in Water Unsaturated Systems

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Bromodeoxyuridine (BrdU) Labeling and Subsequent Fluorescence Activated Cell Sorting for Culture-independent Identification of Dissolved Organic Carbon-degrading Bacterioplankton

Kinetic Visualization of Single-Cell Interspecies Bacterial Interactions

High-Resolution Three-Dimensional Whole-Organ Tomography of Microbial Infections

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

Live Cell Fluorescence Microscopy to Observe Essential Processes During Microbial Cell Growth

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments