8.2: Introduction to Bacterial Identification using Enterotube test - Biology

8.2: Introduction to Bacterial Identification using Enterotube test - Biology

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Learning Outcomes

  • Observe, interpret, and identify bacteria using the Enterotubes/Enteropluri test

Bacterial Identification

The identification of microorganisms is an important part of what many microbiologists do. As you can imagine, clinical microbiologists would need to identify the pathogen that is causing disease in patients. A microbial ecologist is interested in what microbes are contributing to environmental change or they might want to identify new species in the field. Perhaps you are working in a lab and have a contaminant and want to know where it is coming from and what it is. We have already learned some ways to identify microbes, in the last module, you learned about the many biochemical tests available to microbiologists that aid in their identification of bacteria. We’ll continue with some of these biochemical tests, focusing on ones that are capable of combining multiple biochemical tests into one, and the use of selective and differential media to isolate and identify bacteria. Then we’ll move into bacterial genetics and how we can use molecular methods to identify bacteria.


A number of techniques can be used for the identification of specific species and subspecies of Enterobacteriaceae. Speciation is important because it provides data regarding patterns of susceptibility to antimicrobial agents and changes that occur over a period of time. It is also essential for epidemiological studies such as determination of nosocomial infections and their spread.

In an effort to simplify the speciation of the Enterobacteriaceae and reduce the amount of prepared media and incubation space needed by the clinical lab, a number of self-contained multi-test systems have been commercially marketed. Some of these multi-test systems have been combined with a computer-prepared manual to provide identification based on the overall probability of occurrence for each of the biochemical reactions. In this way, a large number of biochemical tests can economically be performed in a short period of time, and the results can be accurately interpreted with relative ease and assurance.

The EnteroPluri-Test /Enterotube is a self-contained, compartmented plastic tube containing 12 different agars (enabling the performance of a total of 15 standard biochemical tests) and an enclosed inoculating wire. After inoculation and incubation, the resulting combination of reactions, together with a Computer Coding and Identification System (CCIS), allows for easy identification. The various biochemical reactions of the EnteroPluri-Test and their correct interpretation are discussed below. Although it is designed to identify members of the bacterial family Enterobacteriaceae, it will sometimes also identify common biotypes of Pseudomonas and other non-fermentative Gram-negative bacilli. It does not identify Pseudomonas aeruginosa.


The EnteroPluri-Test contains 12 different agars that can be used to carry out 15 standard biochemical test. Interpret the results of your EnteroPluri-Test is based on a coding chart included with the test.

The enterotube is a self-contained, compartmented plastic tube containing twelve different media that allow determination of 15 biochemical reactions (glucose, gas production from glucose, lysine decarboxylase, ornithine decarboxylase, hydrogen sulfide (H2S), indole, adonitol, lactose, arabinose, sorbitol, Voges-Proskauer (VP), dulcitol, phenylalanine deaminase (PA), urea, and citrate). The enclosed inoculating wire allows inoculation of all compartments in one step from one or a few single colonies of your unknown microorganism. The resulting combination of enterotube reactions, together with other metabolism tests can help you to identify unknown organisms.

Image 1: Four Enterotubes shown. Uninoculated control tube and 3 tubes inoculated with various bacteria. 24 hr incubation.

Image 2: Enterotube chambers

Interpretation of Enterotube chamber Results:

A. Fermentation of Sugars

Among the common products of carbohydrate breakdown by microorganisms using fermentative pathways are organic acids (acetic, lactic, etc.), alcohols, and gases (carbon dioxide and hydrogen). The types of product formed, and the proportion of each, depends on the species of microorganism as well as the particular carbohydrate being fermented.

The ability to ferment different sugar compounds will be tested by inoculating a single enterotube. The 5 carbohydrates tested in 1 enterotube are: glucose, adonitol, lactose, arabinose, and sorbitol. The formation of acids can be readily detected by including a pH indicator in the microbial growth medium. Acid production will lower the pH of the medium, resulting in a color change of the indicator, creosol red, from red (alkaline) to yellow (acidic).

End products of bacterial fermentation of glucose are either acid, or acid and gas. Gas production will be indicated by the definite and complete separation of the wax overlay from the surface of the glucose chamber. The glucose chamber medium is covered with wax to provide anaerobic conditions to allow detection of gas formation. Fermentation of adonitol, lactose, arabinose, and sorbitol will also result in formation of acidic end products indicated by a change in color of indicator present in the medium from red to yellow. Any sign of yellow is interpreted as a positive reaction; red should be considered negative. Some strains will give slightly variable reactions, refer to chart on page 84-85 for details.

B. Urease

Many bacteria are able to use urea as a nitrogen source by splitting it into ammonia and carbon dioxide through the hydrolysis reaction catalyzed by the enzyme urease:


ǁ urease

NH2 — C—NH2 + H20 ------------------ ► 2NH3 + CO2


The ammonia reacts in solution to form ammonium carbonate, which results in an increase in the pH of the medium. Urease activity is detected by inoculating a medium containing urea and a pH indicator, phenol red (yellow/beige/light amber at acid pH and red-purple at alkaline pH). Initially the urea media chamber is mostly yellow. After incubation, a red-purple color throughout the medium indicates a rapid urea splitter, a positive urease result. No color change, (the agar remains yellow/beige/light amber) is a negative urease result. The urease test is included in the enterotube. Your TA will do the control organisms below:

Escherichia coli does not normally have the enzyme urease. Proteus vulgaris is a rapid urea splitter.

C. Lysine and Ornithine Decarboxylation

Decarboxylase tests are useful for differentiating bacteria. Microorganisms that have the enzyme decarboxylase can remove the carboxyl group from an amino acid. Certain bacteria can decarboxylate the amino acid lysine using the enzyme lysine decarboxylase, which results in the formation of the alkaline end product, cadaverine. Some bacteria can decarboxylate ornithine (a product of arginine hydrolysis) using the enzyme ornithine decarboxylase (ODC), which results in the formation of the alkaline end product putrescine. The presence of these decarboxylation byproducts, cadaverine and putrescine, are indicated by a change in the color of bromcresol purple, the pH indicator in the enterotube medium, from pale yellow (acidic) to purple (alkaline). The medium is covered with wax to provide anaerobic conditions. Any degree of purple should be interpreted as a positive reaction. The medium remains yellow, a negative reaction, if decarboxylation of lysine or ornithine does not occur. These tests are included in the enterotube

Image 3 Lysine and ornithine decarboxylation reactions. The enzymatic reaction catalyzed by ornithine decarboxylase.The pyridoxal phosphate (PLP)-dependent ODC enzyme catalyzes decarboxylation of ornithine and produces putrescine.

D. Voges-Proskauer

Different bacteria convert dextrose and glucose to pyruvate using different metabolic pathways. Some of these pathways produce unstable acidic products which quickly convert to neutral compounds. Some organisms use the butylene glycol pathway, which produces neutral end products, including acetylmethylcarbinol (acetoin) and 2,3-butanediol. The Voges-Proskauer (VP) test detects organisms that utilize the butylene glycol pathway and produce acetoin during glucose metabolism. After inoculation and incubation of the enterotube, the production of acetoin is detected using Barritt’s reagent (potassium hydroxide, and alpha-naphthol) in the VP chamber. The acetoin product is oxidized in the presence of KOH to diacetyl. The diacetyl then reacts to produce a red color. The presence of acetoin, a positive result, is indicated by the development of a red color within 20 minutes, a negative reaction will appear colorless or light amber. The use of Barritt’s reagent in the VP chamber will be performed after all the results are read for the other chambers in the enterotube.

E. Citrate Utilization

This test detects those organisms which are capable of utilizing citrate, in the form of its sodium salt, as the sole source of carbon. Organisms capable of utilizing citrate produce alkaline metabolites which change the color of the indicator from green (acidic) to deep blue (alkaline). Any degree of blue should be considered positive. Certain microorganisms will not always produce the ideal “strong” positive color change. Lighter shades of the same basic color should be considered positive here. The citrate utilization is tested in the enterotube.

How to Inoculate & Interpret an Enterotube

Watch video 1: how to inoculate an Enterotube with your bacterial sample

Watch Video 1: how to inoculate an enterotube performed in Microbiology labs at NC State. (5:08) URL:

Watch Video 2: how to interpret Enterotube results

Watch Video 2: how to interpret Enterotube results. Video by Professor B (5:23). URL:

Microbial identification

9.7 Conclusion

This chapter has outlined some of the microbial identification techniques undertaken. The techniques described have been divided between phenotypic and genotypic methods. It is important to note that groupings established by phenetic and phylogenetic systems do not always agree and within each grouping the methodological differences and varying contents of different databases will sometimes lead to conflicting analyses.

It is additionally important to understand that any systems used to identify bacteria, whether phenotypic or genotypic, will have limitations, because no single test methodology will provide results that are 100% accurate.

In terms of selecting between methods, this will depend on costs and resources, the time that the microbiologist is prepared to wait for and what level of identification is required. Some microbiologists are of the view that the only way to characterize a microorganism correctly is through a “polyphasic approach” that is a combination of phenotypic testing methods and genotypic testing methods. This is, however, far too time consuming and too prohibitively expensive for standard laboratories. Most routine testing laboratories select phenotypic test kits and use established contract test facilities where genotypic testing is required.

What is important, when making a selection, is to go back to basics and consider: what is the purpose of the identification? what does the microbiologist need to know? and what does the result tell the microbiologist? These questions can help with selecting and implementing the appropriate microbial identification test.

Methods for the bioinformatic identification of bacterial lipoproteins encoded in the genomes of Gram-positive bacteria

Bacterial lipoproteins are a diverse and functionally important group of proteins that are amenable to bioinformatic analyses because of their unique signal peptide features. Here we have used a dataset of sequences of experimentally verified lipoproteins of Gram-positive bacteria to refine our previously described lipoprotein recognition pattern (G+LPP). Sequenced bacterial genomes can be screened for putative lipoproteins using the G+LPP pattern. The sequences identified can then be validated using online tools for lipoprotein sequence identification. We have used our protein sequence datasets to evaluate six online tools for efficacy of lipoprotein sequence identification. Our analyses demonstrate that LipoP ( performs best individually but that a consensus approach, incorporating outputs from predictors of general signal peptide properties, is most informative.

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The bacterial strains used in this work are listed in Additional file 1. The entire panel consisted of: i) a collection of Bcc strains representative of all eighteen species described up to now (Peeters et al., 2013, Vandamme and Dawyndt, 2011) and with either clinical or environmental origin ii) 3 Cupriavidus and 3 Ralstonia strains iii) a panel of environmental strains belonging to different bacterial genera provided from CRA-ABP and iv) different clinical strains isolated from CF patients

Materials and Methods

PAFC generates ultrasonic waves resulting from absorption of light in particles under flow. 32 These ultrasonic waves are often created by thermoelastic expansion and contraction of an object that absorbed laser light. 33 , 34 In our PAFC setup, a nanosecond laser operating at 532 nm is used to irradiate a sample under flow. The ultrasonic waves are detected by a piezoelectric transducer and recorded onto a computer. Our photoacoustic sensing setup is directly based on our system used to detect circulating melanoma cells in blood. 35 – 37

Bacteriophages have low optical absorbance at 532-nm wavelength. To provide optical contrast to bacteriophage Det7, Direct Red 81 dye, a polysulfated photostable protein dye capable of generating photoacoustic waves after laser irradiation, was attached. Absorption spectra of dyed and undyed bacteriophage Det7 were measured using a Nanodrop 2300. Bacteriophage remains permanently dyed by Direct Red 81, which contains two sulfonic acid groups with pKa values in the negative range, allowing salt bridges to form with basic groups such as lysine and arginine side chains. These salt bridges are more tenacious than some covalent bonds at relatively neutral pH values, and therefore, these bonds are only broken at an elevated pH in high salt concentrations.

Photoacoustic Flow Cytometry

A Nd:YAG laser (Litron Nano, Bozeman, Montana) coupled into a 1000 μ m , 0.39 numerical aperture, optical fiber (Thorlabs, Newton, New Jersey) was used to produce 532-nm laser light with 5-ns pulses. The laser beam energy coupled through the optical fiber was maintained and measured from 1.9 to 2.1 mJ for most detection experiments. Laser energy was increased to 4 mJ for single cell detection experiment. Laser light was directed to a quartz tube (Quartz 10 QZ, Charles Supper, Natick, Massachusetts) with 10 - μ m -thick walls passing through a 3D-printed flow chamber. The 10 - μ m -thick walls allow the propagation of ultrasonic waves, as well as providing an optically transparent pathway for the sample to flow through. Optical fiber was placed 5 mm from the quartz tube to create a detection volume of 0.04 μ l . Laser beam shape was Gaussian and fluence was calculated to be 0.014 mJ / cm 2 .

A 2.25-MHz transducer focused on the quartz sample tube was fitted to the base of the 3D-printed flow chamber (Fig. 2). The internal volume of the chamber was filled with Sonotech LithoClear acoustic gel (NeXT Medical Products Company, Branchburg, New Jersey) to provide a medium for the propagation of acoustic waves. Syringe pumps were used to create an alternating flow of sample and mineral oil equal to 60/min flow rate. The introduction of sample and the immiscible mineral oil induced two-phase flow. 37 Two-phase flow was employed to allow for future collection of the samples for further analysis while eliminating the possibility of samples becoming stuck or delayed inside the tubing. Signals were amplified with a gain of 50 using a Tegam 4040B amplifier (Tegam, Inc., Geneva, Ohio) and sent to a desktop computer running a customized LabView program. This computer also served for system control and data collection. This flow chamber setup served as the excitation and acoustic wave collection device.

The bacterial cell

All living organisms on Earth are made up of one of two basic types of cells: eukaryotic cells, in which the genetic material is enclosed within a nuclear membrane, or prokaryotic cells, in which the genetic material is not separated from the rest of the cell. Traditionally, all prokaryotic cells were called bacteria and were classified in the prokaryotic kingdom Monera. However, their classification as Monera, equivalent in taxonomy to the other kingdoms—Plantae, Animalia, Fungi, and Protista—understated the remarkable genetic and metabolic diversity exhibited by prokaryotic cells relative to eukaryotic cells. In the late 1970s American microbiologist Carl Woese pioneered a major change in classification by placing all organisms into three domains—Eukarya, Bacteria (originally called Eubacteria), and Archaea (originally called Archaebacteria)—to reflect the three ancient lines of evolution. The prokaryotic organisms that were formerly known as bacteria were then divided into two of these domains, Bacteria and Archaea. Bacteria and Archaea are superficially similar for example, they do not have intracellular organelles, and they have circular DNA. However, they are fundamentally distinct, and their separation is based on the genetic evidence for their ancient and separate evolutionary lineages, as well as fundamental differences in their chemistry and physiology. Members of these two prokaryotic domains are as different from one another as they are from eukaryotic cells.

Prokaryotic cells (i.e., Bacteria and Archaea) are fundamentally different from the eukaryotic cells that constitute other forms of life. Prokaryotic cells are defined by a much simpler design than is found in eukaryotic cells. The most-apparent simplification is the lack of intracellular organelles, which are features characteristic of eukaryotic cells. Organelles are discrete membrane-enclosed structures that are contained in the cytoplasm and include the nucleus, where genetic information is retained, copied, and expressed the mitochondria and chloroplasts, where chemical or light energy is converted into metabolic energy the lysosome, where ingested proteins are digested and other nutrients are made available and the endoplasmic reticulum and the Golgi apparatus, where the proteins that are synthesized by and released from the cell are assembled, modified, and exported. All of the activities performed by organelles also take place in bacteria, but they are not carried out by specialized structures. In addition, prokaryotic cells are usually much smaller than eukaryotic cells. The small size, simple design, and broad metabolic capabilities of bacteria allow them to grow and divide very rapidly and to inhabit and flourish in almost any environment.

Prokaryotic and eukaryotic cells differ in many other ways, including lipid composition, structure of key metabolic enzymes, responses to antibiotics and toxins, and the mechanism of expression of genetic information. Eukaryotic organisms contain multiple linear chromosomes with genes that are much larger than they need to be to encode the synthesis of proteins. Substantial portions of the ribonucleic acid (RNA) copy of the genetic information (deoxyribonucleic acid, or DNA) are discarded, and the remaining messenger RNA (mRNA) is substantially modified before it is translated into protein. In contrast, bacteria have one circular chromosome that contains all of their genetic information, and their mRNAs are exact copies of their genes and are not modified.


Here we empirically show that symbiont genome sizes and functional diversity are predicted by the rate of gene flow into and among symbiont populations via horizontal transmission and homologous recombination. Although we have only investigated three independently evolved associations, we see this system serving as a microcosm for marine associations more generally, as many other associations exhibit similar biologies (e.g., intracellular, autotrophic, broadcast-spawning, etc. [36]) and all are governed by the same population genetic principles. Amazingly, we found that symbiont gene flow between hosts is ongoing in one of the most intimate marine associations, the vertically transmitted vesicomyids. These results suggest that there is a range of possible intermediate genome degradation states that can be maintained over millions of years with sufficient recombination. Therefore, symbiont genome evolution following host restriction is not a one-way, inescapable process that ends in an organelle-like state as it is commonly presented [2,5,6]. These results validate long-standing but untested theory and suggest that the diversity of symbioses found to exhibit intermediate rates of horizontal transmission and incomplete genome degradation may be undergoing similar population-level processes.


Bacterial wilt disease in C. maxima in Guangdong, China, is caused by R. solanacearum. All 24 strains isolated from C. maxima in this study belong to phylotype I, race 1 and biovar 3 or 4. These isolates clustered into sequevars 17, 45, and 56. Sequevar 45 was identified in China for the first time in this study, and sequevar 56 is a novel sequevar that has not been described previously. This study is the first report to identify R. solanacearum race 1 infections in C. maxima. Two C. moschata cultivars, Xiangyu1 and Xiangmi, were resistant or moderately resistant to R. solanacearum strain RS378. Our results provide valuable information for the further development of control strategies for C. maxima wilt disease.


The polymicrobial nature of BV necessitates the use of diagnostic tests that are based on combination criteria. Part of the challenge lies in determining which combination criteria are sensitive and specific enough as diagnostic criteria for BV the other challenge is to develop cost-effective diagnostic tests, which could preferably be used at the POC. Although there is a complex interplay between vaginal pH and the concentration of different bacterial species, it is evident that vaginal pH, specifically higher than 4.5, improves the performance of diagnostic tests when combined with other components in CVF.

The efficient and accurate detection of vaginal dysbiosis has always been plagued by factors such as the difference in biomarker levels across populations (e.g., bacterial species) and sample type variations (Kyongo et al., 2015 Masson et al., 2018). Many studies discussed in this review have highlighted the potential of different combination criteria with biomarkers beyond the genetic level to improve BV diagnostics. If the purpose of diagnosis is to treat, the question should be asked whether a perfect equilibrium exists between the VMB and all its related components. That is, does bacterial concentration translate into corresponding levels of metabolites, proteins and inflammatory markers? Nonetheless, the continual reduction in operational costs of high-throughput technologies provide the opportunity to study the vaginal milieu with a systems biology approach on a large scale to map and link potential biomarkers. This review does not necessarily suggest the replacement of diagnostic tools currently available for BV but does highlight the limitations of these tools and calls for the expansion of the BV diagnostics field by exploring the vast array of diagnostic opportunities discussed here.

In many resource-limited settings, however, POC tests for BV are either not available or simply too expensive for routine diagnostic use and healthcare practitioners have to rely on syndromic management of vaginal discharge syndrome. It is therefore imperative that the development and evaluation of new diagnostic tests must include both a cost- and health-benefit analysis in various settings, especially where expensive instrumentation is required. The risk profiles of different populations for adverse sequelae of BV infection, such as increased risk for HIV infection and poor pregnancy outcomes should guide diagnostic test selection. In such at-risk populations, we have to ask the question—what is the cost of cost?

Category: Microorganisms

Bacterial transformation occurs when a bacterial cell takes up foreign DNA and incorporates it into its own DNA. This transformation usually occurs within plasmids, which are small circular DNA molecules separate from its chromosome. There can be 10 to 200 copies of the same plasmid within a cell. These plasmids may replicate when the chromosome does, or they may replicate independently. Each plasmid contains from 1,000 to 200,000 base pairs. Certain plasmids, called R plasmids, carry the gene for resistance to antibiotics such as ampicillin, which is used in this lab.

Plasmids function in transformation in two different ways. They can transfer genes that occur naturally within them, or they can act as vectors for introducing foreign DNA. Restriction enzymes can be used to cut foreign DNA and insert it into the plasmid vectors. The bacteria used in this lab were Escherichia coli (E. coli). It was ideal for this transformation study because it can be easily grown in Luria broth or on agar, and it has a relatively small genome of about five million base pairs.

Transformation is not the only method of DNA transfer within bacteria. Conjugation is a DNA transfer that occurs between two bacterial cells. A bridge is formed between the two cells and genetic information is traded. In transduction, a virus is used to transfer foreign DNA into a bacterial cell.

The transformed E. coli with the ampicillin resistance gene will be able to grow in the ampicillin plates, but the non-transformed E. coli will not.

The materials needed for this lab were 2 sterile test tubes, 500 μL of ice cold 0.05M CaCl2, E. coli bacteria cultures, a sterile inoculating loop, a sterile micropipette, 10 μL of pAMP solution, a timer, ice, a water bath, 500 μL of Luria broth, a spreading rod, 4 plates: 2 ampicillin+ and 2 ampicillin – , and an incubator.

One sterile tube was labeled “+” and the other “-“. A sterile micropipette was used to transfer 250 μL of ice cold 0.05M CaCl2 to each tube. A large colony of E. coli was transferred with an inoculating loop to each tube. The suspension was then mixed by repeatedly drawing and emptying a sterile micropipette. 10μL of pAMP solution was added to the cell suspension in the tube marked “+” and mixed by tapping the tube. Both tubes were immediately put on ice for 15 minutes and then soaked in a 42° C water bath for 90 seconds. The tubes were then returned to ice for another 2 minutes.

After the heat shock, 250 μL of Luria broth were added to each tube. The tubes were mixed by tapping. Two plates of ampicillin + agar were labeled LB/AMP+ and LB/AMP-. The two plates of ampicillin- agar were labeled LB+ and LB-. 100 μL of the cell suspension in the “+” tube were placed on the LB+ and the LB/AMP+ plates. 100μL of the cell suspension in the “-” tube were added to the LB- and the LB/AMP- plates. These were spread with a spreading rod that was sterilized by passing it over a flame after each use. The plates were allowed to sit for several minutes and then incubated over night inverted at 37° C.

1. Compare and contrast the number of colonies on each of the following pairs of plates. What does each pair of results tell you about the experiment?
LB+ and LB- Both of these plates had a lawn of bacteria. This proves that the bacteria are capable of growing on the agar and that there was nothing preventing growth beside the ampicillin.

LB/AMP- and LB/AMP+ The LB/AMP- had no growth, but the LB/AMP+ had small growth. This shows that the bacteria was transformed and developed a resistance to ampicillin.

LB/AMP+ and LB+ The LB/AMP+ had less growth than the LB+. This shows that the transformation was not completely effective and only transformed some of the most competent bacterial cells.

2. Total mass of pAMP used = 0.05 μg

Total volume of cell suspension = 510 μL

Fraction of cell suspension spread on the plates = 0.196

Mass of pAMP in cell suspension = 0.0098

Number of colonies per μg of plasmid = 0.0294

3. What factors might influence the transformation efficiency? Explain the effect of each you mention.
Transformation efficiency could be affected by the size of the colony added to the solution. In a larger colony the efficiency would increase because there would be more receptive cells. Another factor would b the amount of pAMP added. The more pAMP added, the higher the efficiency. The amount of Luria broth added could also affect efficiency. If the amount of Luria broth was increased, the efficiency would decrease.

Error Analysis:
This lab had several steps, each giving the potential for error. All of the measurements had to be precise and accurate, and the heat shock timing was also a very complicated procedure. Error in this lab could have been caused by the concentration of the CaCl2 due to the fact that most of it was frozen.

Discussion and Conclusion:
The bacteria treated with the pAMP solution developed a resistance to ampicillin and were able to grow on the ampicillin+ plate. Those that were not treated with the pAMP were not able to grow on this medium. The plates with no ampicillin served as a control to show how the bacteria would look in normal conditions. Transformation is never fully effective, Only cells that are competent enough are able to take up the foreign DNA. Therefore, the ampicillin + plates showed less growth than the control plate.

Watch the video: Enterotube (November 2022).