Modeling of Fungal Biofilms Using a Rat Central-vein Catheter

SUMMARY

Candida frequently grows as a biofilm, or an adherent community of cells protected from both the host immune system and antimicrobial therapies. Biofilms represent the predominant mode of growth for many clinical infections, including those associated with placement of a medical device. Here we describe a model for Candida biofilm infection of one important clinical niche, a venous catheter. This animal model system incorporates the anatomical site, immune components, and fluid dynamics of a patient venous catheter infection and can be used for study of biofilm formation, drug resistance, and gene expression.

1. INTRODUCTION

The use of biofilm models has fueled investigation of this unique manner of growth and has been vital for characterizing associated phenotypic properties and gene expression patterns (2, 3, 14). Because Candida infection of commonly placed medical devices, such as a dentures, venous catheters, or urinary catheters, involves biofilm growth, models closely mimicking infection at these clinical sites are of interest (5). To date, the in vivo model most commonly used for Candida biofilm study is the venous catheter model described here (1, 6, 12, 15). This model emulates one of the most common clinical biofilm infections and mimics environmental host conditions at this site, including anatomical location, flow conditions, and exposure to host cells, serum proteins, and immune factors. The catheter is secured in the jugular vein without disruption of blood flow and then tunneled subcutaneously and positioned in a wire casing for protection. Inoculation of the catheter occurs 24 h after catheter placement to allow for a conditioning period of host protein deposition on the catheter surface (5, 7, 11). Throughout the experiment, the model uses the clinically relevant anticoagulant, heparin, although other anticoagulants can be utilized.

The rat venous catheter model can be used as a tool to answer a variety of scientific questions and has identified C. albicans biofilms with altered morphology, adhesion, matrix production and drug susceptibility (10, 13, 15). Confocal microscopy and scanning electron microscopy can successfully illustrate intact biofilm architecture, including the fungal cell morphology, presence of extracellular matrix, and the incorporation of host cells (1). Adjusting the duration of biofilm formation from 6 to 72 hours can capture the time course of this process from cell adhesion to development of a multicellular community with both yeast and hyphal fungal cell morphologies, host cell components, extracellular matrix, and open areas or channels. Sonication effectively removes cells for microbiological enumeration or cellular analyses, such as gene expression profiling or cell biology studies (9). Microbiological counts can be used to quantify the viable biofilm mass and are a simple method of measuring the impact of a luminal drug therapy or comparing the difference in viable burden among several genetic strains. In addition, organs and blood from distant sites can be collected for measurement of viable burden and assessment of biofilm dispersion or dissemination of disease. Although vascular catheters may be infected by hematogenous seeding from a distant vascular site, the model has primarily been utilized for study following intraluminal infection. The latter results in more reproducible cell number and biofilm cell mass among experiments.

2. MATERIALS

2.1 Animals

Specific-pathogen-free male Sprague-Dawley rats weighing 350 g (Harlan)

2.2 Medications

1. Heparin sodium for injection 1000 USP unitis/mL (APP Pharmaceuticals)

2. Xylazine (Sigma-Aldrich,)

3. Buprenorphine 0.3 mg/mL (Hospital Pharmacy)

4. Ketamine HCl 500 mg/10mL (Bedford Laboratories)

5. Double Antibiotic Ointment:Bacitracin Zinc and Polymyxin B Sulfate (Fougera)

2.3 Surgical materials

1. Polyethylene tubing with inner diameter 1.14 mm and outer diameter 1.57 mm. (PE 160, Intramedic, Becton Dickinson)

2. Three way large bore stopcock with rotating male luer lock adapter (Baxter Healthcare Corporation)

3. Rodent jacket, rat 250-350g (Braintree Scientific, Inc)

4. Tether, 18′ sewn (Braintree Scientific, Inc)

5. Scrub Care Surgical Scrub Brush-Sponge/Nail Cleaner (catalog Cardinal Health)

6. Polysulfone Button Tether for rats, 0.090 in lumen, 12in (30cm) (sterile) (Instech Solomon)

7. Skin stapler 5.7 mm × 3.9 mm (Ethicon Endo-Surgery)

8. Surgical suture, sterile, non absorbable, Silk black braided 2-0 18″ (3.0 metric, 45 cm) (Ethicon Inc)

9. Surgical dissecting microscope (Stereo Zoom Microscope with fiber optic illuminator control (PZMIII-BS) World Precision Instruments)

10. Sterile syringes (variety of volumes)

11. Surgical attire: sterile surgical gloves, sterile gown, and surgical mask

12. Rodent hair clipper (A5 power pro clipper, Oster) 13. Rat dissecting kit (World Precision Instruments,)

13. Far Infared warming pad 14″ × 14″ (Kent Scientific Corporation)

2.4 Fungal Isolates and media

2.4.1 Media 1

YPD medium supplemented with uridine: 1% yeast extract, 2% bacto peptone, 2% glucose, and uridine 80 μg/mL

2.5 Materials for evaluation of selected endpoints

2.5.1 Microbiologic counts (optional)

1. Sonicating water bath (FS 14 with 40-kHz transducer, Fisher Scientific)

2. Sabouraud dextrose agar (SDA plates: 4% dextrose, 1% peptone 1.5% agar, pH 5.6

3. Tissue homogenizer (Polytron 3100, Brinkman Instruments)

2.5.2 Confocal or fluorescent microscopy (optional)

1. Fluorescent probes

   Calcofluor white or Fluorescent brightener 28 (Sigma-Aldrich)

   FUN1 live dead yeast stain (Molecular Probes, Invitrogen)

   Concavalin A Alexa Fluor 488 (Molecular Probes, Invitrogen)

2. Glass-bottom petri dish (coverslip 1.5, 35-mm disk P325G 1.5-14C, MatTek)

3. Confocal or fluorescent microscope with inverted objective (such as Zeiss Axiovert 200)

2.4.3. Scanning electron microscopy (optional)

1. Glutaraldehyde (25%) (Sigma-Aldrich)

2. Formaldehyde (37%) (Sigma-Aldrich)

3. Phosphate-buffered saline (PBS) (0.15 M NaCl, pH 7.4)

4. Osmium tetroxide (Electron Microscopy Sciences)

5. Critical point drier (Tousimis)

6. Gold sputter coater (Auto Conductavac IV, Seevac Inc.)

7. Ultra smooth carbon adhesive tabs (12 mm, Electron Microscopy Sciences)

8. Aluminum mounts (12.7 mm, Electron Microscopy Sciences)

9. Scanning electron microscope (JSM-6100, JEOL)

2.5.4. Candida biofilm cell nucleic acid collection (optional)

1. AE buffer (50 mM sodium acetate pH 5.2, 10 mM EDTA)

2. Liquid nitrogen

3. Reagents for hot phenol RNA extraction (1)

3. METHODS

3.1 Preparation of catheters

1. Cut polyethylene tubing into 50 cm in length. This catheter length is calculated based on placement in the jugular vein 2 cm above the right atrium, subcutaneous tunneling, and extension though an external protective device to the top of the animal cage where it will be secured for access. The volume of this catheter length is approximately 500 μL. With luer stub and stop cock, the total catheter volume is approximately 700 μL.

2. Sterilize catheters by ethylene oxide gas sterilization as autoclaving may destroy them.

3.2 Preparation of surgical equipment

1. Sterilize surgical equipment, including surgical gowns, drapes, tethers, and surgical tools by autoclave.

2. Use prepackaged, sterilized stopcocks with luer stubs, sutures, and surgical gloves.

3.3 Catheter placement

1. Anesthetize animals by intraperitoneal injection of a mixture of ketamine (80 mg/kg) and xylazine (8 mg/kg). This anesthesia protocol should produce anesthesia for approximately 120 min.

2. Prepare the animal for the surgical procedure by removing hair from the midscapular space, anterior chest, and neck with a rodent clipper (see Note 2). Prepare skin area with an antiseptic surgical scrub brush.

3. Create a sterile field under the surgical microscope by placing the rat in the supine position and preparing the surgical area with sterile drapes. Wear sterile gloves, mask, and gown.

4. Make a vertical incision in skin of the anterior neck just right of midline and use blunt surgical dissection to expose the right jugular vein.

5. To subcutaneously tunnel the catheter, create a second incision at the scruff and use blunt surgical dissection toward the initial surgical incision. Next, tunnel the proximal end of the catheter through this subcutaneous space to the midscapular space and externalize the catheter at the site of the second surgical incision.

6. Stabilize the jugular vein and make a longitudinal incision of a few millimeters to the vein wall using the vannas scissors. Instill heparinized saline (100 units/ml) into the catheter and insert the catheter in the vein (superior vena cava) opening. Advance to a site above the right atrium (approximately 2 cm). If the catheter is appropriately placed, blood should be able to be easily withdrawn. Conversely, if the catheter is in the atrium, it may be difficult to withdraw blood. Secure the catheter to the vein with (2-0) silk ties.

7. Secure the catheter to the subscapular skin scruff via a button using surgical staples (Fig. 1A). Close both incisions with surgical staples and apply antibiotic ointment. Position a tether and rodent jacket on the animal to protect the catheter (Fig. 1B). Secure the distal catheter segment and stopcock above the cage to allow easy access to catheter.

8. Monitor the animal and wrap in a warming pad until it can lift its head and remain sternal.

9. Administer narcotic analgesia with buprenorphine 0.05 mg/kg subcutaneously twice daily for 24h.

10. Allow the catheter to remain in place for 24 h prior to infection to allow for catheter surface conditioning with host proteins (see Note 3).

Fig. 1.

Surgical placement of a rat jugular venous catheter. (A) The catheter is inserted and secured in the jugular vein of an anesthetized animal. (B) The wire casing and rodent jacket protect the catheter and prevent the animal from disrupting the catheter.

3.4 Animal and catheter maintenance

1. Monitor the animals for signs of distress every 8 h through the study. In necessary, consider additional administration of buprenorphine 0.05 mg/kg subcutaneously twice daily for analgesia.

2. The anterior neck incision and the catheter exit site should be examined daily for signs of inflammation or purulence. In our experience with this protocol, superficial infections are uncommon.

3. House animals in an environmentally controlled room with 12-h light-dark cycle and maintain on a standard ad libitum rat diet. Following surgery and for the duration of the experiments, house animals singly in shoe box cages with normal bedding.

3.5 Preparation of inoculum

1. Store fungal strains in 15% (vol/vol) glycerol stock at -80°C. Prior to experiments, maintain strains on YPD medium supplemented with uridine. C. albicans, C. glabrata, and C. parapsilosis have successfully produced biofilms in this model (8).

2. Grow strains in YPD medium supplemented with uridine at 30°C on an orbital shaker set to 200 RPM. Harvest during late logarithmic phase (this time period can vary among strains thus should be determined experimentally). Enumerate the cells by means of hemocytometer count. Adjust the final density to 1 × 10⁶ cells/mL in YPD supplemented with uridine.

3.6 Infection of catheter

1. Instill 700 μL of fungal inoculum in the catheter using a sterile syringe and the stopcock. This volume should fill the catheter lumen (See Notes 4 and 5).

2. Allow the inoculum to dwell for 6 h, then withdraw or flush the catheter volume. Lock the catheter with same volume of sterile heparinized saline (heparin 100 units/mL, 0.15M NaCl).

3.7 Lock treatment of catheter (optional)

1. Prepare antifungal drugs or other agents to be tested in sterile saline (0.15M NaCl).

2. After 24 h of biofilm growth, withdraw or flush the heparinized saline from the catheter.

3. Instill the drug (700 μL) in the catheter with a sterile syringe and lock in place (see Note 6).

3.8 Harvesting the catheter

1. Sacrifice animals by CO₂ asphyxiation. Typical collection times are 24 h after infection or 24 h after treatment administration (see Note 6).

2. Aseptically remove the catheter from the animal. Collect the proximal catheter segment (approximately 8 cm).

3. Gently place the proximal catheter tip (that was inserted in the animal) on sterile gauze. Allow the catheter fluid to drain the length of the catheter by capillary action (see Note 7).

4. Collect the proximal segment of catheter that was inserted in to the animal (approximately 2 cm in length). This segment can be prepared for microbiological enumeration, microscopy, or nucleic acid collection.

3.9 Endpoint determination

3.9.1 Microbiological counts (optional)

1. Place the catheter section in 1 mL sterile saline.

2. Sonicate sample for 10 min and vigorously vortex for 30 seconds.

3. To ascertain the extent of disease dissemination, remove the kidneys or other internal organs from the animal. Place in a suitable volume of saline and homogenize.

4. Plate serial dilutions (1:10) of the catheter fluid and organ material on SDA plates and incubate for 24 h at 30°C.

5. Enumerate fungal colony counts as an estimate of fungal viable burden per organ.

3.9.2 Fluorescent or confocal microscopy (optional)

1. Cut the catheter segment perpendicular to the catheter length with an 11- blade scalpel into multiple 2- to 3-mm-long “doughnut” segments.

2. Stain the catheter segments with fluorescent probes (FUN1 50 μM, Concanavalin Alexa Fluor 488 conjugate 200 mM, or calcofluor white (22.5 μg/ml) at 30°C for 30 min in the dark.

3. Place catheter segments on the coverslip of a glass-bottom petri dish with the cut edge against the coverslip. Image the luminal surface of the catheter by fluorescent or confocal microscopy using the light source and filters appropriate for selected dyes (1).

3.9.3 Scanning electron microscopy (optional)

1. Cut the catheter segment perpendicular to the catheter length with an 11- blade scalpel into multiple 2- to 3-mm-long “doughnut” segments.

2. Place segments in fixative (1% glutaraldehyde, 4% formaldehyde in PBS) for 16 hours at 4°C.

3. Gently remove fixative and add 1 mL PBS for 10 min to wash samples.

4. Place samples in osmium tetroxide (1% in PBS) for 30 min. Osmium tetroxide is toxic and should only be used in a hood with protective gloves, lab coat, and eye wear. Proper disposal is required.

5. Gently remove osmium tetroxide and add 1 mL PBS for 10 min to wash samples.

6. Dehydrate samples by treating samples to a series of ethanol washes (30% for 10 min, 50% for 10 min, 70% for 10 min, 95% for 10 min, and 100% for 10 min).

7. Use critical point drying according to instruction to accomplish final desiccation. Our protocol uses three 10 minute CO₂ soaks prior to achieving critical point.

8. Section catheter segments length wise and mount the specimens on aluminum stubs with the luminal side visible.

9. Coat samples with gold appropriate for scanning electron microscope using a sputter coater. Our protocol coats samples for 2.5 to 3 minutes.

10. Image the luminal surface of catheter samples using scanning electron microscopy (Fig. 2) (1).

Fig. 2.

Scanning electron micrographs of a Candida albicans biofilm on a rat venous catheter. Catheter segments were harvested, processed, and imaged as described in section 3.9.3. The image at 50x magnification (A) shows the biofilm attached to the luminal catheter surface. At 1000x magnification (B), the hyphae, yeast and extracellular matrix of the biofilm can be visualized.

3.9.4 Nucleic acid collection (optional)

1. Flash freeze catheter segments with liquid nitrogen in AE buffer (50 mM sodium acetate pH 5.2, 10 mM EDTA) (see Note 8).

2. Cells from 2-5 catheters which have been in place for 12-24 h may be pooled to yield higher quantities of RNA (see Note 9).

3. Isolate RNA using the hot phenol method (1).