Author Archives: Admin

Fe-EDTA (A&A)

Na2EDTA•2H2O 866.7 mg
FeSO4•7H2O
or
Fe2(SO4)3•nH2O
2.28 mg
KOH 866.7 mg
Milli-Q Water ~100 mL
  • Dissolve KOH in 31 mL Milli-Q
  • Add Na2EDTA•2H2O
  • In separate container, dissolve FeSO4•7H2O or Fe2(SO4)3•nH2O in 60.67 mL Milli-Q
  • Mix both solutions together
  • Bubble filtered air through combined solutions until the color darkens (~2.5 hr)

0.5M EDTA

Na2EDTA- dihydrate 186.1 g
Milli-Q Water ~1 L
NaOH ~20 g
  • Add EDTA into 800 mL Milli-Q water (NOTE: EDTA will not dissolve)
  • Using NaOH pellets, adjust pH to 8.0 (EDTA should now dissolve)
  • Bring volume to 1 L with Milli-Q
  • Autoclave for 30 minutes
  • Store at 4oC

CI Solution (24:1)

Isoamyl Alcohol 10 mL
Chloroform 240 mL
  • Put on gloves, cover forearms, and relocate to fume hood
  • Add both components to a sterile 250 mL glass bottle
  • Mix well
  • Wrap container in foil to protect from light
  • Store at 4oC

ATCC Culture Medium 929

Ammonium Sulfate 1.32 mg
MgSO4•7H2O 380 mg
CaCl2•2H2O 20 mg
MnCl2•4H2O, 50.5 mM solution 20 µL
Na2MoO4•2H2O, 41.3 mM solution 10 µL
CoCl2•6H2O, 1.7 mM solution 5 µL
ZnSO4•7H2O, 34.8 mM solution 10 µL
K2HPO4 87 mg
Phenol red 0.5% 250 µL
Iron Solution 250 µL
Milli-Q Water ~1 L
  • Measure out approximately 300 mL Milli-Q
  • Add all components in order
  • Bring volume up to 1 L with Milli-Q
  • Adjust pH to 7.5 with 0.5M K2CO3
  • Autoclave for 20 minutes

pH: 7.5

ASNIII

NaCl 25 g
MgCl2•6H2O 2 g
KCl 500 mg
NaNO3 750 mg
K2HPO4•3H2O 20 mg
MgSO4•7H2O 3.5 g
CaCl2•2H2O 500 mg
Citric Acid 3 mg
Ferric Ammonium Citrate 3 mg
EDTA (disodium magnesium) 0.5 mg
Na2CO3 20 mg
Trace Metal Solution 1 mL
Milli-Q Water ~1 L
  • Measure out approximately 300 mL Milli-Q
  • Add all components in order
  • Bring volume up to 1 L with Milli-Q
  • Autoclave for 20 minutes

pH: 7.5

Allen and Arnon Medium (Plates)

-Pi stock 25 mL
+Pi stock 6.25 mL
Agar 10 g
Streptomycin Stock (1 mg/mL) 1 mL
Neomycin Stock (25 mg/mL) 1 mL
Milli-Q Water ~1 L
  • Measure out approximately 300 mL Milli-Q
  • Shake -Pi stock
  • Add both stocks to water
  • Bring volume up to 1 L with Milli-Q
  • Check pH. If less than 7.8, make fresh +Pi
  • Add agar
  • Autoclave for 20 minutes
  • Add antibiotics

Allen and Arnon Medium (Liquid)

-Pi stock 6.30 mL
+Pi stock 3.10 mL
Streptomycin Stock (1 mg/mL) 1 mL
Neomycin Stock (25 mg/mL) 1 mL
Milli-Q Water ~1 L
  • Measure out approximately 300 mL Milli-Q
  • Shake -Pi stock
  • Add both stocks to water
  • Bring volume up to 1 L with Milli-Q
  • Check pH. If less than 7.8, make fresh +Pi
  • Autoclave for 20 minutes
  • Add antibiotics

Cutting Bands from a Gel

Cutting the Bands

  • Fill appropriate number of 500 µL PCR tubes with 50 µL of 10 mM Tris buffer
  • Carefully slide the gel onto the wetted transilluminator plate (wetting the edges of the gel is helpful)
  • Wearing protective equipment, turn on the transmitted UV switch
  • Using a sterilized scalpel (cleaned with 70% ethanol), cut along the top and bottom edges of the band
  • Snip the left and right edges of the band with sterilized forceps (cleaned with 70% ethanol)
  • Remove the excised band and place in corresponding PCR tube
  • Repeat as necessary
  • Store tubes at 4oC for three to five days to allow DNA to diffuse into the buffer

DGGE Protocol

Preparation of Acrylamide Solutions

0% Solution

40% acrylamide/bis (37.5:1, BioRad) 22.5mL
50xTAE buffer 3mL
Milli-Q Water 124.5mL
    • Combine first two components
    • Bring volume up to 150mL with Milli-Q
    • Aliquot into three 50 mL centrifuge tubes
    • Store at 4oC

80% Solution

40% acrylamide/bis (37.5:1, BioRad) 22.5mL
50xTAE buffer 3mL
Formamide (freshly deionized) 48mL
Urea 50.4g
Milli-Q Water ~75mL
    • Combine first four components (urea dissolves slowly)
    • Bring volume to 150mL with Milli-Q
    • Aliquot into three 50 mL centrifuge tubes
    • Store at 4oC

Deionization of Formamide

Formamide 70mL
Mixed Bead Resin 1-2 spoonfulls
  • NOTE: Formamide must be kept wrapped in foil throughout its use due to its sensitivity to light
  • Combine formamide and resin
  • Mix on stir plate for 60 minutes
  • Use Whatman filter paper to remove resin from formamide

Cleaning the Glass Plates

  • Remove one set of glass plates (on tall and one short) and two spacers from the soap solution
  • Wash both sides thoroughly with dish soap and rinse with tap water followed by distillied water
  • Wash the gel-side of each plate with the following reagents in specified order:
    • Windex
    • Acetone
    • 70% Ethanol

Plate Assembly

  • Place the spacers along the edges of the tall glass plate (gel-side up), with the notches facing towards the center of the plate
  • Place the short glass plate (gel-side down) onto the spacers. Align the assemblage with all sides flush, making sure spacers extend slightly past the glass plate edges
  • Lightly tighten the supports to either side of the assemblage, ensuring that the arrows on the supports are facing up and towards each other
  • Place the glass plate assemblage onto the positioning area of the already leveled base
  • Release the tension of the supports and position the plates and the spacers so that they are flush against each other. Proper alignment of the plates is essential for the formation of a good seal
  • Tighten the supports to lock the assemblage into place
  • Pull out the knobs on either side of the base and insert the plates into the pouring area, ensuring that the assemblage is flush against the grey strip
  • Push in the knobs and rotate the screws in a counterclockwise direction (as looking straight down) to secure the plates
  • Verify that there is some bowing in the grey strip to ensure a proper seal

Pouring the Gel

  • Make three solutions from the following chart in 15 mL centrifuge tubes (one of these must be the 0% plug)
0% 10% 20% 30% 40% 50% 60% 80%
 0% solution 6 mL 9.625 mL 8.25 mL 6.875 mL 5.5 mL 4.125 mL 2.75 mL 0 mL
80% solution 0 mL 1.375 mL 2.75 mL 4.125 mL 5.5 mL 6.875 mL 8.25 mL 11 mL
APS (10%) 44 µL 52 µL 52 µL 52 µL 52 µL 52 µL 52 µL 52 µL
TEMED 6.0 µL 6.9 µL 6.9 µL 6.9 µL 6.9 µL 6.9 µL 6.9 µL 6.9 µL
  • Set up the puring apparatus, making sure that the valve connecting the two wells of the gradient maker is closed (~30o from horizontal)
  • Turn on stir bar to 6
  • Center the comb in between the plates and place the syringe between the plates and against one of the spacers
  • Thoroughly mix the acrylamide solutions (20% with either 40% or 60% or 80%) with the APS and TEMED
  • As quickly as possible, pour the higher concentration solution into the front chamber (the one closest to the outlet valve) and pour the lower concentration solution into the back chamber
  • Open the valve between the wells on the gradient chamber (bring to horizontal position) and begin the mixing and the pouring of the gel
  • To promote even mixing, reduce the speed of the stir bar as the gel pours (one should just barely be able to see the mixing in the front chamber). If the stirring speed is too great, the higher concentration denaturant will flow back into the other chamber
  • When the wells are approximately 75% empty, thoroughly combine APS, TEMED and the 0% solution. Add this mixture to the rear well as the front one empties
  • Once both wells are empty, clean the pouring apparatus by running Milli-Q water through the system multiple times
  • Allow the gel to sit for at least two hours. If leaving overnight (recommended), cover assemblage with a large ziploc bag and store in a large tupperware container with a beaker full of water (to reduce dessication)

Assembling the D-Code (Gel Running) System

  • Clean off a set of glass plates (dish soap only)
  • Line the space between the plates with Parafilm
  • Lock plates into place with supports, ensuring they are flush
  • Carefully remove the comb from the gel, along with any excess gel from around the wells
  • With the tall glass facing out and short glass facing in, snap both assemblages into the holder
  • Fill D-Code system with approximately 7 L of 1xTAE (140 mL 50xTAE and 6860 mL Milli-Q water) until fill-line is reached
  • Place the holder down into the buffer, ensuring that it is properly in place
  • Place the top of the D-Code system on top of the apparatus. Verify that all electrical connections are created
  • Plug in electrical chord and turn power switch on
  • Set temperature to 65oC and turn on the heater and pump. Make sure that the buffer level rises and remains above the negative electrode wire
  • Allow D-Code to reach 65oC (90 to 120 minutes)

Loading the Gel

  • Turn off power and remove top of D-Code system
  • Prepare samples by combining equal amounts of template (at least 300 ng) with 5x loading buffer in 500 µL PCR tubes
  • Using a P-100 and corresponding gel loading tips, load samples by placing the pipettor tip half-way into the well and expunging sample with extreme care (loading the wells is made easier by avoiding air bubbles at the pipette tip and expunging slowly)
  • Load the standard into every fourth lane
  • Replace top to the D-Cody system, ensuring all electrical connections are made
  • Plug in D-Code and plug it into both the electrical outlet and the BioRad PowerPack300 (with DCODE chip)
  • Turn on D-Code system and set temperature to 60oC
  • Turn on heater
  • Set PowerPack300 (with DCODE chip) to 200V
  • Turn on the pump only after samples are pulled into the gel (approximately 15 minutes)
  • Let gel run for 3.5 to 4 hours at 200V (or 12 hours at 100V)

Staining and Imaging the Gel

  • Drain the excess buffer from the top of the gel and dismantle the apparatus
  • With the upmost of caution, remove the supports and spacers
  • With extreme care, use one of the spacers as a lever to pry off the short glass plate, leaving the gel on the tall glass plate
  • Soak the tall plate (with gel) in an ethidium bromide stain, and place on rocker (slow mode) for 30 minutes
  • Destain in Milli-Q water for 15-20 minutes
  • Thoroughly wet the imager’s glass plate with water
  • Tilt the glass plate onto the wet surface of the imager, allowing the gel to slowly slide off (continually wetting the edges of the gel is helpful)
  • Once imaging completed and bands to be cut are identified, remove the gel by re-wetting the glass plate and carefully sliding the gel onto it
  • Transport gel to transilluminator for band cutting

DGGE Standards

Materials

ND1-1
ATTC 29133
S1006 (diatom)
MoBio Plant DNA Extraction Kit

or

Materials for PCI Method

Procedure

Extract DNA of all samples using either the MoBio Plant Kit or PCI Extraction method

Amplify DNA with cyanobacterial GC-clamped primers using the standard 30 cycle amplification for cyanobacteria

NOTE: These strains have all been shown to give only one band

Hazardous Waste Pickup

Make sure waste is appropriate, as Risk Management will only accept:

  • Chemicals
  • Biowaste
  • Lamps
  • Batteries

Make sure container is appropriate

  • Condense all waste into as few containers as possible
  • All labels on bottles must be crossed out
  • Bottles must have tight, secure lids
  • DO NOT use biohazard bags for any waste that is not comprised of human tissue or human pathogens
  • Verify that bags are free from tears and leaks

Get a Hazardous Waste Tag (two drawers below hotplate in LSE422)

Fill tag out completely and legibly or waste will not be picked up

  • Chemical: list all chemicals in container using written out names (not chemical formulas)
  • ~~NOTE- they will not pick up any unknowns; everything must have a name~~
  • Amount: rough approximations are fine
  • Vol%: if known
  • Generator: Ferran Garia-Pichel
  • Phone: 727-7762
  • Date: date tag filled out
  • Dept: Microbiology
  • Building: LSE
  • Room: 404, 422, or 427
  • Category:
  • Solid/Liquid: this should reflect container’s contents, not actual contaminant
  • pH: if known

Attach tag to container and set in a visible area where it will not be moved

Contact Risk Management via

Be prepared to provide the following information

  • Your name
  • Your phone number
  • Location of the waste (building, room number, and location within that room)
  • Description of the waste
  • Description of the containers (both type and number)
  • Confirmation that the waste has been tagged
  • Any questions regarding either the waste or the filling out of the tag

Expect to wait a few days before waste is picked up. If nobody comes for it, or it needs to get picked up quickly, call daily

Imaging and Quantifying Gels

Staining

  • Inspect stain and destain solutions to make sure that they are fresh and free of broken gel pieces
  • To make new stain :
    • Pour out existing stain into ethidium bromide waste container
    • Rinse plastic with MilliQ water, pouring rinse into waste container
    • Fill plastic with approximately one liter of MilliQ, and add a few drops of ethidium bromide until water is slightly colored
  • To make new destain:
    • Pour out existing destain into ethidium waste container
    • Rinse plastic with MilliQ water, pouring rinse into waste container
    • Fill plastic with appoximately one liter of MilliQ water
  • Soak the gel in ethidium bromide stain for 30 minutes
  • Use spatula to move gel into a Milli-Q water destain, and let sit for 15 minutes

Positioning Gel

  • Turn on FluorS-MultiImager (power switch is located on back-right hand side)
  • Thoroughly wet the imager’s glass plate with water
  • Remove gel from destain, rinse with Milli-Q water to remove any loose pieces of agarose, and move onto the wet surface of the imager
  • Change gloves
  • Wearing clean gloves, sit at computer and click on “Quantity One” icon (a red icon, found most often on the upper right side of the Desktop)
  • Select “fluorese” from “File” menu
  • Click on “Position” button
  • Use the red plus-mark and the blue grid to center and align the gel
  • Click on “Stop” button and close door to FuorS-MultiImager

Creating Image

  • Set exposure time to 10.0 and click on “Preview” button
    • If bands are too faint, increase exposure time and re-preview
    • If gel is too bright, either decrease exposure time or destain longer
  • Click on “Acquire” button
  • Save image by selecting “Save” from “File” menu, and/or print image by going to “Print” (also under the “File” menu) and selecting “Print Image,” then clicking on “Go” button

Quantifying Bands

  • Open “Lane Tools” and “Band Tools” toolbars (16th and 17th icons from left, respectively, on main Quantity One toolbar)
  • Zoom in on gel (7th icon from left, on main toolbar) as much as possible without cutting off any bands
  • Mark all lanes of interest
    • Select “Create Lane” (6th icon from left, on Lane Tools)
    • Position the mouse above the bands in first lane of interest, centering it horizontally
    • Click, hold, and drag the mouse down past the end of the last band within that lane
    • There should now be a red line running through the center of each band within the first lane
    • Click on “Adjust Lane” (7th icon from left, on Lane Tools)
    • Position mouse over least centered section of line
    • Click, hold and drag the mouse to adjust the line as needed
    • Repeat with all relevant lanes
  • Click on “Detect Bands” (1st icon from left, on “Band Tools”), click on “Close” (all bands should now have brackets roughly surrounding them)
  • Click on “Remove Band” (4th icon from left, on Band Tools)
  • Click on a set of brackets that does not correspond to an actual band, click on “Yes,” and repeat for all other incorrect brackets
  • Adjust brackets vertically
    • Click on “Adjust Bands” (3rd icon from left, on “Band Tools”)
    • Click and hold on the upper bracket corresponding to the first band
    • Adjust bracket both visually and by using the pop-up graphs
    • VISUALLY: The bracket should serve as the upper/lower limits to the bands
    • GRAPHS: Each band has its own peak on the graph. The brackets should be set so that only the steepest section of each peak is highlighted in yellow.
    • NOTE: In theory, each one of these approaches should yield the same result. In practice, such is rarely the case, so it is important to rely on a combination of the two.
  • Adjust brackets horizontally
    • Click on “Lane Width” (10th icon from left, on “Lane Tools”)
    • Click on first lane
    • Arbitrarily adjust number and click “OK”
    • Continue this process until brackets serve as borders to the bands
    • If the (red line demarking the) lane is not centered over the bands it will have to be re-aligned, as it will be impossible to properly adjust the brackets otherwise.
    • NOTE: Re-aligning the lanes will erase the brackets. This time, when preparing to “Detect Bands,” change the “Lanes” option from “All” to “One”, and type in the re-adjusted lane’s number, as labeled by Quantity One.
  • Select “Quantity Standards” from “Analysis” menu, and click on “Create New”
  • Click on the button corresponding to “Select Bands”
  • Return to the window with the gel imange and click on bands from lane containing the mass standard
  • Return to the “Quantity Standards” window
  • Set “Interpolation” to “Linear Regression,” and “Extrapolation” to “Yes”
  • Manually alter the “Quantity” column of the table at the bottom of the window to reflect the known values (if four lanes appear, the values are 20, 15, 10, and 6 for each 1µL added)
  • Click on “Show Curve”
  • If any point is very off, remove it from the chart
  • Click on the button corresponding to “Calibrate Gel,” and click on “OK”
  • Select “All Lanes Report” from “Report” menu, and select “Report”
  • Print report if necessary by choosing “Print All” (2nd icon from left, at bottom of window)
  • Save calculations in personal folder by selecting “Save” from “File” menu (calculations and gel image are all saved together)
  • NOTE: Quantity One caculates the concentration (see “Calibrated Quanitity” column in the “Report” printout) in terms of ng/mL
  • Quit program

Cleanup

  • Remove gel from FuorS-MultiImager and either store in Ziplock bag or throw away in ethidium bromide waste container
  • Clean off FluorS-MultiImager with ethanol
  • Remove any broken pieces of gel from stain and destain, place in ethidium bromide waste container
  • Throw gloves away in ethidium bromide waste container

Ordering Information

If you are not the Research Technician

  • Decide what you want
  • Get a catalogue from outside LSE422 and look up pertinent information :
    • Actual and complete item name,
    • Company name, mailing address, and phone number,
    • Catalogue number,
    • Unit sold (eg each, packet of six),
    • Price per unit
  • Neatly put the catalogue back in alphabetical order (ie where you got it)
  • Give this information, along with amount desired to the technician along with any special concerns (eg rush delivery)
  • Wait 7 to 10 days

NOTE: you must provide the aforementioned information to the technician even if you place an order with the credit card

If you are the (acting) Research Technician

  • Make sure you are provided all necessary infomation from the grad student/professor
  • Get the order approved by Dr. Garcia-Pichel, and ask which grant to use for the order
  • Get a green departamental order form from the file cabinet
  • Fill out the form, using completed ones as templates to answer any questions
  • Get Dr. Garica-Pichel to sign it
  • Photocopy the form, and turn the original over to Kristy Bolen (LSE 210)
  • File the photocopy according to grant number under the “incomplete” heading
  • Upon arrival, double check accuracy and condition of item(s)
  • If there are no complications:
    • Write “received okay” on packing slip along with date
    • Photocopy slip
    • Give original slip to K. Bolen
    • Attach photocopy of slip to photocopy of order form
    • File photocopies appropriately (according to year and account number)
  • If there are complications:
    • Leave order as inctact as possible
    • Find order form and double check order
    • Take both packing slip and order form to K. Bolen (LSE 210) and explain the problem
    • From here, everything will be situation-specific
  • It is your responsibility to keep track of orders and make sure they arrive. Many companies will not contact ASU if there is a problem with the order, be it their fault or ours.

PCI Extraction

Reagents and Solutions

TESC Buffer 11 mL/sample
Proteinase K 30 µL/sample
10% SDS 550 µL/sample
PCI ~15 mL/sample
CI ~10 mL/sample
Isopropanol ~6 mL/sample
70% Ethanol (-20oC) ~0.5 mL/sample
10 mM Tris or TE Buffer 250 µL/sample

 

Materials

Scalpel, autoclaved 1
Tweezers, autoclaved 1
Weigh Boats, autoclaved 1 per sample
All-Glass Tissue Grinder or ceramic mortar and pestle, autoclaved 1 per sample
1.5 mL Eppendorf Tubes, autoclaved 1 per sample
100 µL Pipette 1
100 µL Pipette Tips, autoclaved one per sample
1 mL Pipette 1
1 mL Pipette Tips, autoclaved many
5 mL PCI-Pipette (located in the hood in LSE404) 1
5 mL Pipette Tips, autoclaved many
Water Baths one a 37oC
one at 50oC
one at 65oC
50 mL Plastic Centrifuge Tubes with Screw Caps four per sample

Sample Preparation

  • Autoclave everything
  • Using sterile tweezers, scalpel and weigh boat, weigh out approximately 50-300 mg wet mass of sample (flame sterilize tweezers and scalpel between samples)
  • Transfer sample into all-glass tissue grinder (potter), add 1 mL TESC buffer and homogenize
  • Using 1 mL pipette, transfer homogenate to 50 mL centrifuge tube
  • Rinse potter with 4 mL TESC buffer and combine with above homogenate
  • Rinse potter with 1 mL TESC buffer and combine with above homogenate
  • Freeze at -80oC (~20 minutes), thaw at room temperature for ~5 minutes, thaw completely in 65oC water bath (~10 minutes)
  • Repeat above sequence five times (for a total of six freeze-thaws)

Extraction

  • Add 30 µL Proteinase K solution and 550 µL 10% SDS solution to sample
  • Gently mix by inverting capped tube several times
  • Incubate at 50oC for 40 minutes, gently mixing every 10 minutes
  • Add 5 mL PCGently mix by inverting capped tube several times
  • Centrifuge at 4,100 rpm for 10 minutes
  • Carefully transfer aqueous (ie top) layer into a new 50 mL centrifuge tube
  • Add 5 mL TESC buffer to original tube
  • Gently mix by inverting capped tube several times
  • Centrifuge at 4,100 rpm for 10 mintues
  • Combine aqueous (ie top) layer with that from above
  • Add one volume (~10mL) PCI to combined aqueous layers
  • Gently mix by inverting capped tube several times
  • Incubate at 65oC for 10 minutes, gently mixing every five minutes
  • Centrifuge at 4,100 rpm for 12 minutes
  • Transfer aqueous (ie top) layer into a new 50 mL centrifuge tube
  • Add one volume (~10mL) CI
  • Gently mix by inverting capped tube several times
  • Centrifuge at 4,100 rpm for 12 minutes
  • Transfer aqueous layer to new 50 mL centrifuge tube
  • Centrifuge at 4,100 rpm for 12 minutes
  • Remove any remaining PCI/CI from tube
  • Gently mix by inverting capped tube several times
  • Centrifuge at 4,000 rpm for 8 minutes

Culmination

  • Precipitate nucleic acids with 0.6 volume (~6 mL) isopropanol for at least 30 minutes at room temperature, or overnight at 4oC
  • Alternatively, add 0.04 volume (~400 µL) 5 M NaCl and mix, add 2 volumes (~20 mL) cold (-20oC) 100% ethanol and mix, precipitate for 60 minutes in an ice bath
  • Centrifuge at 4,100 rpm for 60 minutes at 4oC
  • Decant supernatant
  • Wash pellet with cold (-20oC) 70% ethanol, pouring it slowly along the tube’s wall. If pellet dislodges, re-centrifuge at 4,100 rpm for 20 minutes at 4oC
  • Dry pellet for 1-3 hours in vacuum
  • Add 250µL 10 mM Tris (or TE) buffer to tube
  • Incubate at 37oC for at least 30 minutes to allow for dissolution
  • Transfer re-suspended pellet into a 1.5 mL Eppendorf tube
  • Store at -20oC

PCR

Materials

  • PCR tubes (one per sample, two for controls, and one extra)
  • Two 2 mL microcentrifuge tubes
  • Template
  • E. coli DNA (or other positive control)
  • Primers, forward and reverse
  • 10% BSA (LF)
  • 10x Ex Taq buffer
  • dNTP mixture
  • 5x Eppendorf Enhancer
  • Milli-Q water, autoclaved
  • Ex Taq DNA polymerase
  • Assorted pipettors and tips

Procedure

  • Open all PCR and microcentrifuge tubes and place in racks found in the PCR hood, along with all pipettors and tips (note: all pipettors should already be in hood)
  • Close the side panels and set the UV light timer for 20 minutes
  • Turn on UV light and blower for the flow hood
  • Gather all reagents and samples, placing them in an ice bath
  • Enhancer should be placed in a warm water bath to dissolve precipitate
  • Enter relevant data into the “Master Mix” calculation spreadsheet; make a print out
  • In PCR hood, prepare master mix in one 2 mL microcentrifuge tube
  • Aliquot master mix into PCR tubes
  • Transfer PCR tubes to ice bath and move to flow hood
  • Add template to each reaction tube, E. coli (or equivalent) to positive control, and PCR water to negative control
  • Place caped and labeled tubes in PCR thermocycler, then close lid
  • Turn on thermocycler, select desired PCR cycle, and begin
  • Pause thermocycler at very end of second cycle
  • Quickly add Taq polymerase to each tube (note: Do not expel polymerase from tip; it will flow out automatically. Do not allow any air bubbles to form in PCR tube.)
  • Close thermocycler lid and press “resume”
  • Store tubes at 4oC

PCR Cycles

Cyanobacterial Primers (cyano)

First Cycle (run once)

  • 95oC for 5 minutes
  • 80oC for 1 minute
  • With about 3 seconds remaining, pause thermocycler and add Taq

Second Cycle (run 30 times)

  • 94oC for 1 minute
  • 60oC for 1 minute
  • 72oC for 1 minute

Third Cycle (run once)

  • 72oC for 9 minutes
  • 4oC hold

Eubacterial Primers (eubac td)

First Cycle (run once)

  • 95oC for 5 minutes
  • 80oC for 1 minute
  • With about 3 seconds remaining, pause thermocycler and add Taq
  • 65oC for 1 minute
  • 75oC for 3 minutes

Second Cycle, Touchdown PCR (run 20 times)

  • 94oC for 1 minute
  • 65oC for 1 minute
  • 75oC for 3 minute
  • NOTE: this is a touchdown cycle; the temperature for the second step must be lowered by 1oC every second cycle (ending on 55oC).

Third Cycle (run 9 times)

  • 94oC for 1 minute
  • 55oC for 1 minute
  • 72oC for 3 minute

Fourth Cycle (run once)

  • 72oC for 7 minutes
  • 4oC hold

Converting Sequencing Files

Converting from PC to Mac

  • Save/transfer all PC files into a single folder on the computer
  • Open ConvProg (desktop alias named “Sequence Converter”)
  • A standard folder select dialog box will appear
  • Navigate to the folder containing all files to be converted (note: do not open said folder, simply select it)
  • Click the “choose” button

Agarose Gel Protocol

Assembling Electrophoresis Cell

  • Gather all parts. These should be located on the shelf above the pipette rack, but can often be found in/near the sinks or in the 4oC : Base, Gel-Casting Gates (two), Gel Tray,Comb(s),Safety Lid with Electrical Cables
  • Place the base on an open part of the counter
  • If base is not level, adjust Leveling Feet accordingly
  • Insert the Gel-Casting Gates into their corresponding grooves
  • Place the Gel Tray between the Gel-Casting Gates
  • NOTE: there is a nice diagram in the BioRad Life Science Research Products catalog on page 255

Preparing the Gel

  • Combine either 40 mL buffer with 400 mg agarose in an Erlenmeyer flask for a small gel or at least 100 mL buffer with 1.0 g agarose, to create a 1% solution
  • Use 1x TAE as your buffer if your samples are DNA extracts, and 1x TBE if your samples are PCR amplified DNA
  • Plug up flask with a paper towel
  • Microwave flask for roughly one to two minutes for a small gel and two to three minutes for a large gel to dissolve agarose, stopping microwave and swirling flask every time it shows signs of boiling
  • Once completely dissolved, cool solution under running water until the flask can be held without burning skin
  • Double check that Gel-Casting Gates are securely in place and pour agarose solution onto Gel Tray
  • For small gels, pour all 40 mL; for large gels, fill between two and three ticks (these are on the sides of the base)
  • Rinse out Erlenmeyer flask, fill with water, and set aside
  • Pop any bubbles in gel with a syringe needle
  • Insert appropriate Comb(s) into the appropriate Comb Slots
  • Allow gel to solidify
  • Thoroughly clean out Erlenmeyer flask
  • Gel can now be saved for later use. Place in Ziplock bag and store at 4oC

Loading Samples

  • Remove Comb(s) and Gel-Casting Gates
  • Wipe gel off Comb(s) and Gel-Casting Gates with paper towel to remove any agarose residue
  • Fill base with same buffer as above to cover gel
  • On a strip of Parafilm, measure out 5 µL sample and 2 µL Nucleic Acid Sample Loading Buffer, 5x
  • Mix sample and loading buffer by slowly sucking in and out of pipettor tip
  • Load samples into wells
  • Never use outside lanes
  • Dip pipette tip into buffer on side with red Bananna Plug/Electrode Cassette before loading well to remove any excess dye
  • Carefully lower tip half-way into well, being careful not to touch gel
  • Load well slowly to ensuring none of the sample escapes into the buffer
  • Raise pipette tip out of buffer before releasing thumb (very important)
  • Load 5 µL Precision Molecular Mass Standard into one well and 1 µL into another

Running Gel

  • Place Safety Lid on Base, matching up Electrical Cables to Bananna Plugs/Electrode Cassettes according to color
  • Plug Electrical Cables into power source (PowerPack 300 or EC105)
  • Turn on power source, set voltage to 110V for small gels and 120V for large gels, and press the “running man” button to begin (verify that the gel is running by looking for a stream of bubbles originating at the black Bananna Plug/Electrode Cassette)
  • Run small gels for approximately 20 minutes and large ones until the dye is 75% across the gel
  • Turn off and unplug power source, and disconnect Electrical Cables
  • Remove Safety Lid and Comb(s)
  • Remove Gel Tray with gel on it (gel can either be imaged or stored in a Ziplock bag at 4oC)
  • Pour buffer out of Base into a labeled container and store at 4oC for later use
  • Clean out Base, Comb(s), and Gel Tray

Imaging Gel and Quantifying Bands

  • Follow standard procedure

Soil microbial ecology

Niche differentiation of ammonia-oxidizing microbial communities and their function in soil

Soil archaea and bacteria are known to oxidize ammonia to nitrite in a key pathway of nitrification. The nitrogen (N) cycle may be affected by N inputs from natural (e.g. mineralization) or anthropogenic (e.g. atmospheric deposition) sources. Most research has shown that N enrichment alters ammonia-oxidizing microorganisms, increasing ammonia oxidation (AO) rates and abundance of ammonia-oxidizing putative enzymes. However, the archaea and bacteria, and subgroups within each microbial group, may respond uniquely to available NH4+ concentrations and to changes in the environment. We ask, what are the dynamics that control ammonia-oxidizing communities and their effects on ecosystem processes? We used soils from N fertilized (NH4NO3 added at 60 kg N ha-1 yr-1 since 2005) and unfertilized Sonoran Desert soils near Phoenix, Arizona, to measure AO using the nitrite-accumulation method. To test for effects of patch type in aridlands, soils were also collected away from plants and under the canopy of creosote shrubs. In the lab, we measured potential rates using shaken-slurries and actual net rates using static incubations. Rates were measured under a range of starting NH4+ concentrations to develop a response-curve of AO kinetics. Additionally, ammonia-oxidizers were quantified using real-time PCR and identified to the species level using clone libraries and pyrosequencing (data processed with Qiime).

Long-term N fertilization increased rates of potential and actual AO in soil away from and under plants. Based on molecular analyses, N fertilization increased diversity and absolute number of ammonia-oxidizers in the total community. Additionally, one archaeon population makes up 74-95% of all the ammonia-oxidizers across treatments and patch types in these desert soils. Interestingly, inspection at a fine level of resolution within the archaea and bacteria reveal that many individual populations either increase or decrease, exhibiting niche separation through a community shift. Furthermore, the rate of AO per copy number of ammonia-oxidizing cells (i.e. AO efficiency), increases with N fertilization. These results suggest that environmental N addition in aridland soils alters ammonia-oxidizing communities at the genetic level and elevates nutrient cycling rates at the ecosystem scale.

Link to 2013 CAP-LTER Symposium Poster by Yevgeniy Marusenko, Ferran Garcia-Pichel, and Sharon J. Hall

Litter decomposition in arid systems

Litter decomposition is a key pathway in the global carbon (C) cycle, releasing more C into the atmosphere annually than fossil fuel combustion. While litter decay in mesic systems is reasonably well predicted by empirical models based on climatic and litter chemistry factors, this is not the case in arid systems. Specifically, mass loss in arid systems is faster than predicted and decay patterns are near linear rather than exponential. Recent research has revealed that breakdown of litter by solar radiation (photodegradation) can be a significant driver of mass loss in arid systems, although the relative strength of this driver appears quite variable. The UV component of sunlight appears to be the most effective waveband in driving mass loss, and lignin appears to be the main target. We propose that the optical properties of leaf litter vary substantially among different plant growth forms, and that this has a large influence on the effectiveness of photodegradation. We suspect that the surface UV-screening of leaf litter is greatest in evergreens, intermediate in grasses, and least in forbs. Screening effectiveness is important because it dictates radiation fluxes inside litter, such as in vascular tissue where the majority of lignin resides. We also suspect that effective surface screening persists much longer in evergreen litter than in grasses or forbs, because of the high concentrations of wall-bound screening compounds in evergreens that are not be readily lost during decay. Hence, photodegradation may become a stronger driver of decay much earlier in the decomposition of forb and grass litter than evergreen litter. We contend that these changes in the potential effectiveness of photodegradation during decomposition may explain the relatively linear litter decay patterns, as well as the variable effectiveness of photodegradation, found in arid systems.

Microbialites

Microbialites

Stromatolites and Oncolites from Desert Springs

The Cuatro Ciénegas Basin (Coahuila, Mexico) is a complex karstic system in which the underlying Cretaceous limestone, dolomites and gypsum formations are actively dissolved by an aquifer of distant origin. This results in the formation of innumerable springs, surface and underwater streams, caves and sinkholes pozas, which are famous for their beauty and the biological diversity they harbor. Within the frame of a large multidisplinary effort funded by NASA’s National Astrobiology Institute, scientist at ASU are looking at the food-web stocihiometry, biosignatures, grazer interactions, and microbial populations of these springs.

Cyanobacteria are often dominant primary producers in calcareous freshwater springs. In most cases, they occur as sessile, benthic or epiphytic dwellers, and are also typically associated with the precipitation of the microcrystalline calcite, that often results in the formation of macroscopic stromatolitic structures and rolling oncolites. These systems allow us to study the interactions between microbial metabolism and carbonate precipiation, in a manner that may help us understand present and past microbialites.

Jet-Suspended, Calcite Ballasted Waterwarts

While studying the Cuatro Ciénegas microbialites, we noted planktonic populations of marble-sized colonies of blue-green algae developing at Escobedo’s Warm Spring, a sheltered, small, fast-flowing spring. There, cm-sized waterwarts were kept in suspension by the upwelling waters of a central 6-m deep well. Waterwarts were built by an Aphanothecelike unicellular cyanobacterium and supported a community of epiphytes that included filamentous cyanobacteria and diatoms, but were free of heterotrophic bacteria on the inside. Waterwarts contained orderly arrangements of mineral crystallites, made up of microcrystalline low-magnesium calcite, with high levels of Strontium and Sulfur. An analysis of the hydrological properties of the spring well and the waterwarts demonstrated that both, large colony size and the presence of controlled amounts of mineral ballast, are required to prevent the population from being washed out of the well. The mechanisms by which controlled nucleation of extracellular calcite is achieved remain to be explored.

 

Mircrobial Biosignatures

Microbial Biosignatures

What are biosignatures?

Any evidence of life

Why are they useful for?

Look for past life, look for life on other planets, to understand life.

Why ‘microbial’?

From all the life we know, prokaryotes (archea and bacteria) are the most widespread organisms, in time and space. They are the first organisms that populated the Earth, and would likely be the first dwellers on other planets where life may have developed. They have also the widest limits of environmental tolerance (temperature, pH, radiation, desiccation, etc.), the widest variability in metabolic strategies, and they occupy all the known ecological niches. Additionally, they have been the dominant form of life for about 70% of the geologic time. Thus, they should be the starting point for the search of ancient life on Earth and beyond.

Evidence of the existence and activity of microbes in the fossil record consists primarily on stromatolites, microfossils, and biomolecules, whose antiquity can go far back to the Archean (~3.5 Ga). These microbial signatures can be traced almost countinually over time since their appearance on Earth, but the older they are, the hardest to prove them biogenic and the easiest to confuse them with abiogenic structures. Hence, because their biogenicity becomes less evident, not one, but many biosignatures should be retrieved from the study objects, and these biosignatures should converge into a unique conclusion supporting a life-originating process. Otherwise, the object could be discarded as life-related.

What kind of fossil biosignatures exist?

Life can be manifested in several ways, and thus traced using:

  • Biomarkers : chemical compounds produced inside cells
  • Biominerals : minerals produced by their influence on the environment
  • Bioisotopes : isotopes derived from metabolic activity
  • Ichnofossils, microbialites or biofabrics : sedimentary structures biologically originated
  • Microfossils : any cell remains

The fact is:

  • We need to understand life to understand biosignatures
  • Biosignatures must prove biogenicity or non abiogenic origin
  • Lots of biotic and abiotic processes, their products and effects are still unknown
  • Morphology may be valid, but more independent biosignatures confirming biogenicity are encouraged
  • As technology, knowledge and interdisciplinarity advance, more biosignatures will be able to detect, and more things known about them.

Biosignatues of terrestrial microorganisms

Biological Soil Crusts(BSCs) are organo-sedimentary structures formed by microbes(mainly dyanobacteria, but also fungi, byrophytes, and algae) that cement topsoil sediments forming a crust.

crusts diagram

Modern BSC are important because they:

  • increase/decrease runoff of rain water
  • hydrate topsoil for a longer time
  • water from below escape at lower rates
  • prevent/decrease soil erosion
  • contribute to C, N and other nutrients in soils
  • enhance seed anchoring and germination
  • are distributed all over the world(principally in arid to semiarid areas

Hypothesis

Because cyanobacteria, the main builders of biological soil crusts, are a very old taxa (~3,500 Ma), very well adapted to UV-light and desiccation conditions, they were probably dwellers on the early Earth’s land. They likely colonized and formed crusts in incipient soils as they do today. But yet, there is no evidence for that. To prove that they existed in the Precambrian would have deep evolutionary and ecological implications.

How to distinguish BSC in rocks?

Many other structures that resemble BSCs are found in the rock record (varves, mud cracks, cohesive sand layers, etc.). Thus, unique features of modern BSCs must be recognized in the rocks.

crusts poster

We study sedimentological, chemical and biological features of a variety of BSCs and other structures. The architecture of modern BSCs, its variability within depositional settings, and the morphological changes under diagenetic processes may provide useful elements to distinguish crust-like structures from the rock record.

We found that the crust tends to be enriched in metals as compared to the soil underneath the crust and the uncrusted soils. Minerals are slightly different also, and characteristic sedimentary structures develop only when microbes are present. So, microbes are leaving behind their signature in many of the aspects we study. The metabolic diversity should be further studied so to understand more on the interactions between microbes and particles.

So far, we can say that soil type, water regime, local temperature and wind may influence the interaction between microbes and sediments, which together determine the structure and constitution of BSCs. Differences among BSCs from different sedimentary environments comprise crust thickness, topology, porosity, and layering. Layers of fine sediment, formed by microbial action, can be preserved and recognized after compaction. These features may be useful as indicators of microbially-produced crusts and may be identifiable in the rock record.

Given the great diversity of abiotic sedimentary structures found in nature, individual features alone cannot be counted as biosignatures. Several biosignatures (biominerals, bioisotopes, biomarkers, microfossils, etc.) must be considered when looking for BSC in the rock record. Finally, a definition of BSCs should encompass an array of characteristics that distinguish them from other crust-like structures. This work is a preliminary approach to that goal.

Hypersaline Mats

Hypersaline Communities from Baja California

As part of a large scale, multidisciplinary effort to understand the complex interactions between microbial community structure and emergent ecosystem properties we are studying hypersaline microbial mats from Guerrero Negro in Baja California (México) as models of microbial ecosystems.

These photosynthetic communities are benthic, laminated aquatic biofilms consisting of highly structured, and dynamic communities of microorganisms. Hypersaline microbial mats harbor a large variety of organisms able to tolerate and thrive under environmental extreme conditions such as high salinity, light exposure and gases such as hydrogen sulfide and hydrogen.

We are particularly interested in explaining the distribution and abundance of these microorganisms in time and space in terms of environmental gradients and interactions. These microbial mats represent the modern analogs for what must have been the major type of ecosystem on Earth for much of its early history. Understanding their functioning gives us the key to understanding the past features of Earth’s biogechemistry.

Sunscreens

Sunscreens

bacteria-sunscreen

Solar ultra violet radiation or UVR (wavelengths shorter than 400 nm) is associated with biological deleterious effects in living organisms. Among these, some cyanobacteria must thrive in habitats exposed to high doses of UVR such as soil and rock surfaces, and thus have the ability to synthesize and accumulate UV-sunscreens. Sunscreens serve as passive preventative mechanisms that allow the organism to stop UVR before it reaches the cellular machinery, DNA, or produces reactive oxygen species.

The indole-alkaloid, scytonemin, found exclusively among cyanobacteria, is one such sunscreen. It is a brownish-yellow, lipid-soluble pigment located in the extracellular matrix of the cells. The production of scytonemin is induced by UV-A (315-400 nm) and the conjugated double-bond distribution allows for the molecule to absorb strongly in that range (with a maximum of ~384 nm). Scytonemin also has potential in biomedical applications because of its strong anti-proliferative and anti-inflammatory activity.

Another class of sunscreens found in cyanobacteria are the mycosporine-like amino acids (MAAs), water soluble, colorless products that absorb and are induced by UV-B (280-315 nm).

Our lab is currently focusing on the molecular genetics of scytonemin and MAAs biosynthesis.

The model organism that we use to study sunscreen biosynthesis is the filamentous heterocystous cyanobacterium Nostoc punctiforme ATCC 29133/PCC 73102 (from the order Nostocales).
N. punctiforme was originally isolated from the symbiotic association with the gymnosperm cycad Macrozamia sp. and it is one of the few cyanobacteria that can grow heterotrophically. In addition, the fact that N. punctiforme is amenable to genetic manipulation (by electroporation or conjugation) and its genome is fully sequenced (US Department of Energy’s Joint Genome Institute (JGI) database), make it a good model for our work.

Using N. punctiforme, we have been able to obtain a scytonemin-deficient mutant by random transposon insertion into a putative gene. The genomic region of mutation has been identified and is currently being studied for its significance in the biosynthesis of scytonemin, as well as its presence in other scytonemin-producing cyanobacteria.

Biological Soil Crusts

Definition: Surface-bound assemblages of microorganisms consolidate soils into mm to cm-thick crusts that occur on arid lands wherever the lack of water restricts the settlement and development of plant cover. To know more about Biological Soil Crusts, please explore the further sub-sections.

Introduction :

Biological soil crusts (BSCs) are also known as cryptogamic or cryptobiotic microbial communities. They are complex microbial communities dominated by cyanobacteria as primary producers that build crust on the top layer of arid lands soils.

Facts numbers:

  • 35% of the total Earth’s continental surface is covered by arid lands, BSC usually cover these areas
  • 30 to 350 kg C ha-1 is the annual range of carbon input in BSC
  • 1 to 100 kg N ha-1 is the annual range of nitrogen fixation in BSC
  • 4th is the position of Microcoleus vaginatus in the World ranking of the most abundant cyanobacteria
  • 54 x 1012 g of Carbon is the biomass of microbial primary producers in BSC

In spite of their geographic extent and ecological importance, many aspects of the biology of BSCs remain unknown; this is why we are studying them!

BSC formation, story of a very slow process

1) Crusting is initiated by growth of filamentous cyanobacteria (e.g. Microcoleus sp.) during episodic events of available moisture.
2) As they grow, these cyanobacteria produce a high amount of slime (extra-polymeric substances) that traps mineral particles.
3) This process result in the formation of a pioneer light-crust
4) Once the crust is stabilized other microbes colonize the crust. For instance other cyanobacteria (e.g. Nostoc sp.) forms black colonies on the top of the crust
5) Later on, other organisms, such as lichens, eukaryotic microalgae, and mosses, may be integrated as dwellers of the crust.

Greening deserts: the incredible ability of Microcoleus sp.

When BSC get wet they rapidly turn green.
This is due to Microcoleus sp. that moves to the surface of the crust once exposed to water. These bacteria are doing oxygenic photosynthesis and carry green-blue photosynthetic pigments; this is why the surface of the soil turns green when they move there.
The answer of Microcoleus sp to wetting event corresponds to a quick metabolic shift that allow the turn ON of metabolism pathways.
Once the crust get dry, Microcoleus sp go back inside the crust, a few millimeters below the surface. Microcoleus sp filament can survive for long periods in this hostile environment ( dry, with low light and low nutrients input) through days, months or years…until the next wetting event.
The behavior of Microcoleus sp. and more generally of the whole crust microbial communities through these wetting-drying cycles is studied in collaboration with research groups from LBL and JGI.

Objective :

We are interested in the description of how the community switch ON when the crust get wet and then switch OFF and get prepared to long periods of dessication when the crust dry.

Approach:

To describe this process we are using a combination of cultivation, direct DNA/RNA sequencing, metabolites identification, in silico modelling and imagery based techniques.
If you want to know more about this project, please contact us!
You can also visit the webpage of our collaborators in Laurence Berkeley laboratory.

References :

1) F Garcia-Pichel, J Belnap, S Neuer, F. Schanz – Algological Studies , vol. 109, 2003 Estimates of global cyanobacterial biomass and its distribution
2) F Garcia-Pichel, O Pringault – Nature, 2001 Microbiology: Cyanobacteria track water in desert soils
3) F Garcia‐Pichel, J Belnap – Journal of Phycology, 1996 MICROENVIRONMENTS AND MICROSCALE PRODUCTIVITY OF CYANOBACTERIAL DESERT CRUSTS

Destruction and recovery

Human disturbances in Biological Soil Crust (BSC) often create severe environmental problems. The dust storms or “Haboobs” that have been increasingly striking Phoenix area in the recent years are a good local example. Biological Soil Crusts are responsible for maintaining soil cohesion and stability in arid areas. They also improve the soil fertility and play an important role in the germination, growth and survival of native species of plants.
Our main research objective is to facilitate the recovery of degraded arid and semi-arid lands through the restoration of the biological soil crusts. This project is developing laboratory methodologies and establishing a nursery for testing the inoculation techniques and biocrust formation. The inoculation techniques will also be tested and monitored in the field, operating in a degraded biocrust from the Department of Defense Training Areas. Special attention will be paid on whether biocrust restoration may facilitate the function of native plant vs. colonization of exotic plants.

References:

Garcia-Pichel, F., Lopez-Cores, A., and Nubel, U. (2001) Phylogenetic and morphological diversity of cyanobacteria in soil desert crusts from the Colorado Plateau. Applied and Environmental Microbiology 67: 1902-1920.

Belnap, J., and Lange, O.L. (2001) Biological soil crusts: Structure, function, and management. Spirnger-Verlag. Berlin, 479 p.

Zaady, E., Gutterman, Y., and Boeken, B. (1997) The germination of mucilaginous seed of Plantago coronopus, Reboudia pinnata, and Carrichtera annua on cyanobacterial soil crust from the Nagev Desert. Plant Soil 190: 247-252

Garcia-Pichel, F., and Belnap, J. (1996) Microenvironments and microscale productivity of cyanobacterial desert crust. Journal of Phycology 32: 774-782.

Image gallery

Euendolithic Cyanobacteria

Insights into the world of euendolithic cyanobacteria.

There are several classes of endolithic organisms: euendoliths actively carve out or bore into mineral material, chasmoendoliths live in the crevices and cracks of mineral material, cryptoendoliths live within structural cavities of porous rock inluding previously excavated now vacant euendolithic dwellings.

Our research focuses on euendolithic organisms which bore into calcium carbonate containing minerals. Carbonate minerals like calcite or dolomite, and other carbonates like dead coral, carbonate sand, marble sculptures, fountains or even concrete represent some of the preferred substrates for euendolithic microbes. The reasoning behind the evolution of this particular lifestyle is poorly understood but we believe that by choosing this niche they improve their chances of survival, and consequently play both friend and foe in the environment taking an important role in the rock cycle (via bioerosive forces), and sometimes accelerating the deterioration time of monuments made of carbonates.

Autofluorescence test

Cyanobacteria are photoautotrophs which utilize the sun’s radiant energy to ultimately produce glucose which is then respired to produce ATP. The stars of our research are the true endoliths or “euendoliths” that actually dissolve the rock matrix, boring into the substrate and making a tunnel where the cells spend their life. This biogenic destruction of carbonates contributes to erosion and the transformation of the mineral matrix into microcrystalline carbonate or micrite. The process is not always negative; in some microbial communities like in the case of stromatolites the micritization actually contributes to the growth of these structures by cementing sediments together, providing support. This micrite layer often encases the organism which can lead to microfossil formation. The fascinating thing about these boring microbes is that we do not really know how they do it.

test BD-TH

Some researchers have suggested various mechanisms including mechanical destruction, symbiosis with heterotrophs, dissolution by acid secretion, and active transport of metal ions by ATP driven pumps, the later being our proposed mechanism of action.
Our current model organism, Mastigocoleus testarum strain BC008, is a marine cyanobacteria isolated from the coast of Cabo Rojo, a coastal town on the island of Puerto Rico. This particular microbe dissolves calcium carbonate by a mechanism that is not well understood and is the topic of our groups research. We hypothesize boring is mediated by a series of ATP-driven calcium pumps as well as calcium channels (García-Pichel, 2006). Our experimental approach includes using calcium-sensitive fluorescent dyes like Calcium Green 5N and confocal microscopy to try to image the dissolution of calcite (crystalline calcium carbonate) in situ. Other techniques include the use of calcium pump blockers to evaluate the effect on the boring activity, as well as measuring boring activity in minerals other than calcite. We are also interested in the genetics behind carbonate boring and will be utilizing high throughput RNA-seq in an attempt to identify transcripts putatively involved in carbonate boring.

Mechanisms of Carbonate Dissolution by Cyanobacteria.

Among the many interactions between biology and geology, the formation and subsequent destruction of carbonates stands as one of the most conspicuous and widespread. At the microscale one can go from calcification in biofilms to the accretion of coralline atolls. Historically biogenic carbonate precipitation has received the most attention, but its dissolution can also be mediated by organisms, and by microorganisms in particular. Fungi, microalgae and cyanobacteria that actively bore into calcareous substrates have been known for more than a century, and have been leaving fossils and trace fossils since the Precambrian. These organisms are fairly ubiquitious in both marine and fresh water ecosystems as well as in many terrestrial envronments. The mechanisms by which many of these euendolithic organisms bore into carbonate substrates has been studied predominately in fungi and microalgae. The boring mechanistics of euendolithic cyanobacteria are largely unknown but our lab has recently developed a model system to study cyanobacterial euendolithic carbonate boring in the laboratory.

Colonies of Hyella sp. growing on marine carbonates.

Colonies of Hyella sp. growing on marine carbonates.

Biogenic carbonate precipitation has received the most attention, but its dissolution can also be mediated by organisms, and by microorganisms in particular. Fungi, microalgae and cyanobacteria that actively bore into calcareous substrates have been known for more than a century, and have been leaving fossils and trace fossils since the Precambrian. These boring microorganisms are centrally implicated in a variety of geological phenomena, ranging from the erosive morphogenesis of coastal limestones, the destruction of coral reefs, the reworking of carbonaceous sands and the cementation of stromatolites. But for all their significance, the mechanism by which they can excavate carbonates in a controlled manner remains to be studied. The most common hypothesis as to their action mechanisms has been that they dissolve limestone by excretion of acids. However, we contend that, in the case of photosynthetic organisms like cyanobacteria, their activity constitutes an apparent paradox, since the dissolution of carbonates runs contrary to the well-known geomicrobial effects of oxygenic photosynthetic metabolism, which will tend to make the surrounding medium alkaline and therefore promote calcification, not carbonate dissolution. We will test three alternative models than can explain cyanobacterial boring and still be consistent with thermodynamic, physiological and mineralogical constraints. Two models are based on the separation of photosynthetic and respiratory activities (either temporally or spatially). The third model is based on localized and directed cellular calcium transport. We will undertake a three-tiered experimental approach using cultivated microorganisms and well characterized mineral substrates that should offer evidence regarding the validity of each of these models.

We are currently using: a) long-term monitoring of the rates of growth and boring with manipulations of various environmental parameters, b) short- term studies of microscale mass transfer in actively boring systems, including the effects of specific inhibitors, using microsensors, and c) advance microscopy studies of active mineral /microbe systems that offer both visual and micro-chemical information: laser scanning confocal microscopy using extracellular and intracellular calcium fluorophores, fluorescent localization, and secondary ion mass spectroscopy (SIMS).

References:

García-Pichel, F. Ramirez-Reinat, E. Gao, Q., 2010. Microbial excavation of solid carbonates powered by P-type ATPase-mediated transcellular Ca2+ transport. Proc. Natl. Acad. Sci. U.S.A. 107:21749-21754.

García-Pichel, F., 2006. Plausible mechanisms for the boring on carbonates by microbial phototrophs. Sedimentary Geology 185 (2006) 205-213

See also an example of past microborers.

Hydrogenases

Interest in the generation of renewable fuels has gained momentum in the last decades in the face of global warming associated with the continued use of fossil fuels, and because of the finite nature of their reserves.

Biohydrogen production from photosynthetic organisms constitutes a conceptually promising avenue in renewable bioenergy, because it would couple directly solar radiant energy, essentially inexhaustible, to the generation of clean, carbon-neutral biofuels, particularly if water-splitting (oxygenic) phototrophs were used.

Cyanobacteria, the only group of oxygenic phototrophs among the bacteria, have been regarded as good models for research and eventual application in this area for several reasons: they are capable of growth with minimal nutritional requirements, they are demonstrable producers of hydrogen gas under certain physiological conditions, and some can be genetically modified with ease. Among cyanobacteria, three different enzymes participate in hydrogen metabolism nitrogenase, and two types of Ni-Fe hydrogenases (uptake and bidirectional).

hydrogenase

 

In principle, production of biohydrogen based on nitrogenase systems requires significant modifications of the enzyme or cumbersome growth conditions in order to promote proton reduction and prevent N2 reduction. The hydrogen produced by nitrogenase is often recycled back into metabolic reducing equivalents by means of the uptake hydrogenase. Under physiological conditions, and as the name suggests, the latter enzyme can only consume, rather than produce H2, and so does not constitute a viable platform for biohydrogen production; in fact, it needs to be inactivated to improve yields of nitrogenase-based hydrogen production. Certain cyanobacteria, however, host a bidirectional hydrogenase that can catalyze both the production and the uptake of hydrogen under physiological conditions. One of the major disadvantages for sustained H2 production via the bidirectional hydrogenase is the easy reversal of the reaction direction.

Therefore, our lab focuses on surveying newly isolated cultures from diverse environments for the presence of the bidirectional hydrogenase gene and a concurrent quantitative comparison of their hydrogenase activities under non nitrogen-fixing conditions. We target cyanobacteria from terrestrial environments, since no bidirectional hydrogenase genes originating in these environments were known from public databases, suggesting that they may have been differentially under-sampled. Marine microbial intertidal mats were also of special interest since a high flux of hydrogen had been reported from these cyanobacterial mats. To this we add a survey of freshwater plankton, a habitat well known to harbor cyanobacteria with bidirectional hydrogenases and hydrogen producing capabilities.

Hydrogenase-2

Microbial Adaptation to Arid Conditions

While the desert seems to be an inhospitable place, in reality it contains a multitude of diverse microorganisms. Many of these microorganisms live in the crust, the top several mm to cm of soil. In this environment, organisms survive high levels of UV irradiation with very little water. I am interested in ‘how microorganisms can live in such an environment?’.

Much of what is known about desiccation resistance comes from the Class Deinococci. Deinococcus radiodurans, the most famous member of the Class, was initially isolated from a can of spoiled meat that had been irradiated for sterilization. D. radiodurans has been shown to withstand 500,000 rads of radiation and still maintain the viability of some cells (Mattimore & Battista, 1996. J. Bacteriol. 178:633-637). PFGE was used to analyze the effect of irradiation on the genome – the results showed the breakdown of the full length genome, approximately 3 Mbp in length, into small fragments estimated to be 50 kbp. Over time, the wild-type irradiated organisms were shown to rebuild a full-length functional, stable, genome from these fragments (Harris et al., 2004. PLoS Biol. 2: e304:1629-1639). Our overriding question is: does the mechanism that allows D. radiodurans to survive irradiation occur in organisms present in the desert crust?