13.15: Bacterial Identification Methods

Bacterial Smear Preparation

Cells, such as bacteria, are colorless (and small) which means they are difficult to observe microscopically. The cytoplasm is essentially transparent and specimens need to contrast with the background of the field. It is then necessary to use a stain (i.e., a dye) to be able to visualize them. The staining methods outlined in this page are used to observe and study the size, morphology, arrangement, and cellular structures of bacteria. The quality of the staining depends on the preparation of a suitable smear. Its thickness or concentration can affect the stain, cells should also adhere to the slide to resist one or more washings.

A bacterial smear, also known as an emulsion, is a thin layer of bacteria spread on the surface of a clean microscope slide. Proper smear preparation is essential to ensure effective staining and accurate observation of bacterial cells. The thickness of the smear is a critical factor. A smear that is too thick contains a high concentration of cells, which reduces light transmission through the specimen and makes individual cells difficult to observe. Excessive thickness can also interfere with proper staining by trapping the stain, potentially leading to false-positive results. A thin bacterial smear does not have enough cells making it difficult to find cells, the staining might not be clear and the cells too interspersed to assess their arrangement. The bacterial load in a smear depends on the source of the sample. In liquid media, bacteria are suspended throughout the broth, typically resulting in a more diluted sample. In contrast, bacteria grown on solid media form dense colonies on the agar surface, leading to a higher concentration of cells.

Another factor to consider when preparing a smear is adherence. Adherence of the cells to the slide is critical, as they must withstand the application of stains and the wash steps. Before fixing it is essential to air-dry the smear. This allows excess moisture to evaporate, preparing the sample for proper heat-fixation. Doing so will prevent the sample from boiling when passed over the heat, which results in the breakage of the cells. Once air-dried, the slide is quickly passed over the flame 3 to 4 times, without exposing the cells directly to the flame. Heat-fixation not only firmly attaches the cells but it also kills the cells, making them safer to handle. It also prevents autolysis, the self digestion of cells, by denaturing enzymes. Finally, heat-fixation coagulates cytoplasmic proteins, improving cell visibility.

The two protocols below outline how to prepare a smear from a liquid broth and from a solid medium. In addition, Figure \(\PageIndex{1}\) illustrates the major steps of preparing a bacterial smear.

Lab Protocol: Smear Preparation from a Broth

1. Obtain a clean microscope slide and mark an area in the center of the slide with a grease/wax pencil.
2. Open the tube containing the inoculated liquid broth and pass the mouth through the flame several times.
3. Sterilize an inoculating loop by passing it several times through the flame. Allow it to cool down.
4. Aseptically collect a loopful of broth and deposit it into the marked area on the slide. Carefully disperse the broth in this area using the loop.
5. Repeat step 4 two more times.
6. Air dry the slide. Do not use the flame to dry the smear.
7. Once dry, heat-fix the slide by running it through the flame back and forth 2 times.
8. Allow the slide to cool.

Lab Protocol: Smear Preparation from Agar

1. Obtain a clean microscope slide and mark an area in the center of the slide with a grease/wax pencil.
2. Sterilize an inoculating loop by passing it several times through the flame. Allow it to cool down.
3. Aseptically transfer 3 to 4 loopful of water into the marked area on the slide.
4. Sterilize the inoculating loop by passing it several times through the flame. Allow it to cool down.
5. Aseptically lift a bacterial colony from the plate with the inoculation loop.
6. Disperse the specimen in the water drop on the glass slide.
7. Air dry the slide. Do not use the flame to dry the smear.
8. Once dry, heat-fix the slide by running it through the flame back and forth 2 times.
9. Allow the slide to cool.

Common Staining Methods

There are several different stains that can be used to stain bacterial smears. Some of the most common are:

• Methylene blue (simple stain)
• Gram staining (differential stain)
• Acid Fast staining (differential stain)
• Endospore staining (differential stain)

Lab Protocol: Simple Staining with Methylene Blue

The protocol below outlines how to stain bacterial cells with methylene blue, a positively-charged dye that binds to the negatively charged molecules of a cell, causing them to stain blue. This protocol is an example of a simple staining method (Figure \(\PageIndex{1}\), in which only a primary stain is used.

1. Prepare a bacterial smear from a liquid broth or a colony.
2. Stain the smear with a 1% methylene blue solution for 1 minute.
3. Gently rinse the stain away with distilled water.
4. Place the slide between two sheets of bibulous paper (or paper towel) and carefully blot dry.
5. Observe the slide.

Figure \(\PageIndex{1}\): Simple staining. Steps 1 to 4 (top box) show the production of a bacterial smear. Step 1: An area is outlined on a clean microscope slide with a wax pencil (outlined in blue). Step 2: A sample of bacterial broth culture, or bacterial colony, is obtained using a sterile inoculating loop. Step 3: The sample is applied to the area on the slide and then the smear is air dried. Steps 2 and 3 are repeated until sufficient bacteria is added to the slide. Step 4: The sample is quickly heat fixed and then allowed to cool completely before stained. Steps 5 through 7 (bottom box) show the simple staining procedure with methylene blue. Step 5: The primary stain (i.e., 1% methylene blue) is added to the area and allowed to sit on the cells for 1 minute. The dye is rinsed well with water. Step 6: The slide is gently dried by placing between two sheets of bibulous paper. Step 7: The stained bacterial cells (rods and circles) are observed under the microscope. (Simple Staining by Patricia Zuk, CC BY 4.0; figure created in BioRender. Zuk, P. (2025))

Methylene Blue Staining Reagents

1% Methylene Blue

• add 1.0 g methylene blue powder to 80 mL distilled water
• mix well
• adjust volume to 100 mL with distilled water
• store at room temperature

Gram Staining

Gram staining is an important staining method universally used in clinical microbiology. It differentiates between two main types of bacteria cells, Gram-positive or Gram-negative, so named based on whether they stain with the crystal violet component of the Gram stain. Gram staining is an example of a differential staining method, meaning it uses more than one dye. To learn more about Gram staining, go to Chapter 7.2: Microbial Identification.

Lab Protocol: Gram Staining

1. Prepare a bacterial smear from a liquid broth or a colony.
2. Stain the smear with a 2% crystal violet solution for 1 minute.
3. Gently rinse the stain away with distilled water.
4. Add the Gram's iodine stain to the slide and let sit for 1 minute.
5. Gently rinse the stain away with distilled water.
6. Decolorize the slide using 95% ethanol for ten to fifteen seconds.
   • alternate decolorizer solutions include acetone or an acetone:ethanol solution
7. Rinse the slide of decolorizer with distilled water.
8. Apply the 0.25% safranin O counterstain for 1 minute.
9. Gently rinse the stain away with distilled water.
10. Pat dry with bibulous paper.
11. Observe the slide.

Figure \(\PageIndex{2}\): Gram Staining. Step 1: A 2% crystal violet solution is added to a bacterial smear and allowed to stain the sample for 1 minute. The slide is rinsed of stain with distilled water. At the end of this step, both Gram positive (+) and Gram negative (-) cells will stain purple. Step 2: An iodine solution is added to the slide for 1 minute to act as a mordant. The slide is rinsed with distilled water. At the end of this step, both Gram positive (+) and Gram negative (-) cells be stained purple. Step 3: The sample is quickly decolorized of crystal violet stain for 10 to 15 seconds using 95% ethanol. The slide is rinsed with distilled water. After this step, only the Gram (+) cells will be stained purple (purple circles). Gram (-) cells will be decolorized and will be colorless (gray rods). Step 4: A 0.25% safranin O solution is added for 1 minute to counterstain the sample. The slide is rinsed of counterstain with distilled water. At the end of this step, the Gram positive (+) cells will be purple (purple circles) and the Gram negative (-) cells will be pink (pink rods). (Gram Staining by Patricia Zuk, CC BY 4.0; figure created in BioRender. Zuk, P. (2025))

Gram Staining Reagents

2% Crystal Violet

• dissolve 2 g of crystal violet in 20 mL of 95% ethanol
• add 80 mL of distilled water and mix well
• store at room temperature in a light-protected bottle

Gram's Iodine

• dissolve 2.0 g of KI (potassium iodide) powder in 100 mL of distilled water
• add 1.0 g of iodine crystals and mix until fully dissolved
• adjust volume to 300 mL with distilled water
• store at room temperature in a light-protected bottle

0.25% Safranin O

• dissolve 0.25 g of safranin O powder in 10 mL of 95% ethanol
• add 90 mL of distilled water and mix well
• store at room temperature in a light-protected bottle

Acetone-Ethanol Decolorizer

• combine 50 mL of acetone with 50 mL of 95% ethanol
• store at room temperature

The quality of the Gram stain depends on several key factors. One of the most critical is the age of the bacterial culture. Fresh cultures typically have intact, rigid cell walls that respond well to Gram staining, while older cultures may contain dying cells with compromised cell wall integrity, leading to inconsistent or weak staining results. The quality of the smear itself also plays a significant role. Thick smears tend to retain more dye, and areas of high cell density may appear overly saturated with crystal violet, potentially obscuring the counterstain and leading to false interpretation. Another major variable is the decolorization step, which is often the most sensitive part of the process. Both over-decolorization and under-decolorization can occur easily, even within the recommended time window of a few seconds. The optimal duration of this step depends on the smear’s thickness—dense smears typically require more time, while thinner smears need less. Optimal decolorizing time will require empirical determination and experience.

Acid-Fast Staining

An acid-fast stain is a differential staining technique that identifies organisms that have a waxy, lipid-rich cell wall (rich in mycolic acid) that is resistant to standard stains. Under treatment with acid-alcohol, the organisms will retain certain dyes. Acid-fast staining is typically used to stain Mycobacterium species like M. tuberculosis. Acid-fast staining can be performed using a "hot" method (i.e., the Ziehl–Neelsen Method) that uses heat to help the primary stain penetrate the cells, and a "cold" method (i.e., the Kinyoun Method) in which the heating step is eliminated but uses higher concentrations of the primary stain. To learn more about acid-fast staining, go to Chapter 7.2: Microbial Identification.

Lab Protocol: Ziehl–Neelsen Acid-Fast Staining

1. Prepare a bacterial smear from a liquid broth or a colony.
2. Cover the smear with Ziehl-Neeson carbol fuchsin stain.
3. Gently heat the slide until steam appears from the smear. Keep steaming the slide for 5 minutes.
   • Do not boil the bacterial smear or allow it to dry out
   • Reapply stain if the smear dries out
4. Let the slide cool for 1 minute.
5. Gently rinse the stain away with distilled water.
6. Apply the acid-alcohol stain until the runoff is clear (about 10 to 30 seconds)
7. Immediately rinse the slide with distilled water.
8. Cover the smear with 1% methylene blue for 1 minute.
9. Gently rinse the stain away with distilled water.
10. Pat dry with bibulous paper.
11. Observe the slide.

Figure \(\PageIndex{4}\): Ziehl-Neeson Acid-Fast Staining. Step 1: A carbol fuschin solution is added to a bacterial smear. At the end of this step both acid-fast positive (+) and acid-fast negative (-) cells are stained pink. Step 2: The slide is gently heated for approximately 5 minutes. The slide is allowed to cool and rinsed with distilled water. At the end of this step, both acid-fast (+) and acid-fast (-) cells are stained pink. Step 3: The sample is quickly decolorized of crystal violet stain for 10 to 30 seconds with a 3% acid-alcohol solution. The slide is immediately rinsed with distilled water. After this step, only the acid-fast (+) cells will be stained pink (pink rods). Acid-fast (-) cells will be decolorized and will be colorless (gray circles). Step 4: A 1% methylene blue solution is added for 1 minute to counterstain the sample. The slide is rinsed of counterstain with distilled water. At the end of this step, the Acid-fast (+) cells will be pink (pink rods) and the acid-fast negative (-) cells will be blue (blue rods). (Ziehl-Neeson Staining by Patricia Zuk, CC BY 4.0; figure created in BioRender. Zuk, P. (2025))

Ziehl-Neeson Acid-Fast Staining Reagents

Ziehl-Neeson Carbol Fuschin

• dissolve 5.0 g of phenol crystals (i.e., carbolic acid) in 90 mL of warm distilled water
• dissolve 0.3 g of basic fuchsin power in 10 mL of 95% ethanol
• combine the two solutions and mix thoroughly.
• store in a dark, tightly sealed bottle at room temperature
• use caution - phenol is toxic and corrosive

3% Acid-Alcohol Decolorizer

• slowly add 3.0 mL of concentrated HCl to 97 mL of 95% ethanol and mix well
• store in a labeled, sealed bottle
• caution - solution is flammable and corrosive

1% Methylene Blue

• dissolve 1.0 g of methylene blue powder in 100 mL distilled water
• filter if necessary
• store at room temperature

Lab Protocol: Kinyoun Acid-Fast Staining

1. Prepare a bacterial smear from a liquid broth or a colony.
2. Cover the smear with Kinyoun carbol fuchsin stain and let stand for 5 to 10 minutes.
3. Gently rinse the stain away with distilled water.
4. Apply the acid-alcohol stain until the runoff is clear (about 10 to 30 seconds)
5. Immediately rinse the slide with distilled water.
6. Cover the smear with 1% methylene blue for 1 minute.
7. Gently rinse the stain away with distilled water.
8. Pat dry with bibulous paper.
9. Observe the slide.

Kinyoun Acid-Fast Staining Reagents

Kinyoun Carbol Fuschin

• dissolve 8.0 g of phenol crystals (i.e., carbolic acid) in 80 mL of warm distilled water
• dissolve 4.0 g of basic fuchsin power in 20 mL of 95% ethanol
• combine the two solutions and mix thoroughly.
• store in a dark, tightly sealed bottle at room temperature
• use caution - phenol is toxic and corrosive

Acid-Alcohol Decolorizer

• slowly add 3.0 mL of concentrated HCl to 97 mL of 95% ethanol and mix well
• store in a labeled, sealed bottle
• caution - decolorizer solution is flammable and corrosive

1% Methylene Blue

• dissolve 1.0 g of methylene blue powder in 100 mL distilled water
• filter if necessary
• store at room temperature

Endospore Staining

The endospore stain is a differential staining technique that detects bacterial endospores, which are highly resistant, dormant structures formed by some bacterial genera like Bacillus and Clostridium. The most common method is the Schaeffer–Fulton method. In this method, heat, in the form of steam, is used to help the primary stain, malachite green, penetrate the spore coat. Water is used to rinse the primary stain away and decolorize the specimen. This method will distinguish endospores from the vegetative cells of the bacterial specimen. To learn more about endospore staining, go to Chapter 7.2: Microbial Identification.

Lab Protocol: Schaeffer-Fulton Endospore Staining

1. Prepare a bacterial smear from a liquid broth or a colony.
2. Cover the smear with 5% malachite green.
3. Place the slide over boiling water or a steam bath and let stand for 5–10 minutes.
   • Keep the stain from drying out by adding more if needed.
4. Let the slide cool for 1–2 minutes.
5. Gently rinse the stain away with distilled water.
   • this step will also decolorize the sample
6. Cover the smear with safranin O for 30–60 seconds.
7. Gently rinse the stain away with distilled water.
8. Pat dry with bibulous paper.
9. Observe the slide.

Lab Protocol: Quick Endospore Staining

1. Prepare a bacterial smear from a liquid broth or a colony by heat-fixing 20 times over the flame
2. Cover the smear with 5% malachite green and let stand for 15 minutes.
   • Keep the stain from drying out by adding more if needed.
3. Gently rinse the stain away with distilled water.
   • this step will also decolorize the sample
4. Cover the smear with safranin O for 30–60 seconds.
5. Gently rinse the stain away with distilled water.
6. Pat dry with bibulous paper.
7. Observe the slide.

Figure \(\PageIndex{5}\): Endospore staining. Step 1: A 1% malachite green solution is added to a bacterial smear. Step 2: The slide is gently heated over boiling water for 5 to 10 minutes and then allowed to cool for 1 to 2 minutes. At the end of this step, both the bacterial cells (rods) and spores (circles) are stained green. Step 3: The sample is quickly decolorized of the stain for 10 to 30 seconds with distilled water. After this step, only the spores will be stained green (green circles) and the bacterial cells will be colorless (grey rods). Step 4: A 1% safranin O solution is added for 1 minute to counterstain the sample. The slide is rinsed of counterstain with distilled water. At the end of this step, the bacterial cells will be pink (pink rods) and the spores will be green (green rods). (Spore Staining by Patricia Zuk, CC BY 4.0; figure created in BioRender. Zuk, P. (2025))

Endospore Staining Reagents

5% Malachite Green

• add 5.0 g malachite green (or malachite oxide) powder in 100 mL distilled water
• use gentle heat to fully dissolve the powder
• filter the solution if needed
• store at room temperature in a light-protected bottle

1% Safranin O

• dissolve 1.0 g of safranin O powder in 10 mL of 95% ethanol
• add 90 mL of distilled water and mix well
• store at room temperature in a light-protected bottle

Carbohydrate Fermentation Testing

A carbohydrate fermentation test (i.e., a fermentation test) is a type of biochemical test that is used to determine whether a bacteria (or yeast) can ferment specific sugars. Bacteria, or yeast, are grown in a Carbohydrate Broth containing nutrients, a specific sugar for fermentation, and a pH indicator. Common sugars used in this test include glucose, lactose, sucrose, maltose, mannitol, galactose, cellulose, and starch. To detect gas production, a small, thin glass tube, called a Durham tube, is inverted into the broth. The presence of air within the Durham tube at the end of the incubation period will be a positive indication of gas production. The use of a pH indicator will confirm the production of acids. The most common pH indicator used is Phenol Red which will turn the broth yellow with the production of acids. Other pH indicators include Andrade's indicator (colorless to pink) and Bromothymol Blue (blue to yellow). To learn more about carbohydrate testing, go to Chapter 7.2: Microbial Identification.

Lab Protocol: Carbohydrate Fermentation

1. Prepare a Carbohydrate Broth.
2. Aliquot 5 to 10 mL of broth into sterile test tubes.
3. Invert and submerge a sterile Durham tube into each test tube, ensuring that it fills completely and that there is no air in the Durham tube.
4. Loosely cap the tubes.
5. Autoclave to sterilize. Let cool.
6. Label each tube with the organism's name and type of sugar being used for fermentation.
7. Aseptically inoculate each tube with the test organism using a sterile loop.
8. Incubate the tubes at 37°C for 24 to 48 hours.
9. Observe and record results.

Fermentation Reagents

Carbohydrate Broth with 0.18% Phenol Red

• to 90 mL of distilled water, add the following
  • 1.0 g Peptone
  • 0.5 g NaCl
  • 1.8 mL of a 1% Phenol Red stock solution
  • 1.0 g of the specific carbohydrate to be tested
• mix until all components are dissolved (heat to boiling if required)
• adjust pH to 7.0
• adjust volume to 100 mL with distilled water
• autoclave at 121°C for 15 minutes to sterilize
• let cool
• store at 4°C

API Testing

In microbiology, API strips refers to the Analytical Profile Index (API), a standardized identification system used to identify bacteria and yeast based on their biochemical activities. Each API strip is composed of multiple chambers, called cupules, that are specific to a metabolic function or enzyme. A small aliquot of cellular suspension is added to each chamber and then incubated. The results are then analyzed and recorded. Based on the results, the microbe can be identified and classified. To read more about API testing, go to Chapter 7.2: Microbial Identification.

Lab Protocol: API Test Strips

1. Select a bacterial colony and prepare a bacterial suspension in sterile saline or sterile distilled water.
   • a suspension of yeast cells can also be prepared
2. Fill each cupule of the test strip with the cell suspension using a Pasteur pipette.
   • anaerobic cupules should be covered with mineral oil
3. Place the strip in its incubation tray and add a few drops of water to maintain humidity.
4. Incubate at 37°C for 18–24 hours
   • incubation times will vary by test panel and organism being tested
5. Observe color changes in each well
   • some cupules will require the addition of another reagents
6. Record the pattern of positive/negative results.