Abstract
Infectious diseases pose a formidable global challenge, compounded by the emergence of antimicrobial resistance. Consequently, researchers are actively exploring novel antimicrobial compounds as potential solutions. This endeavor underscores the pivotal role of methods employed for screening and evaluating antimicrobial activity—a critical step in discovery and characterization of antimicrobial agents. While traditional techniques such as well-diffusion, disk-diffusion, and broth-dilution are commonly utilized in antimicrobial assays, they may encounter limitations concerning reproducibility and speed. Additionally, a diverse array of antimicrobial assays including cross-streaking, poisoned-food, co-culture, time-kill kinetics, resazurin assay, bioautography, etc., are routinely employed in antimicrobial evaluations. Advanced techniques such as flow-cytometry, impedance analysis, and bioluminescent technique may offer rapid and sensitive results, providing deeper insights into the impact of antimicrobials on cellular integrity. However, their higher cost and limited accessibility in certain laboratory settings may present challenges. This article provides a comprehensive overview of assays designed to characterize antimicrobial activity, elucidating their underlying principles, protocols, advantages, and limitations. The primary objective is to enhance understanding of the methodologies designed for evaluating antimicrobial agents in our relentless battle against infectious diseases. By selecting the appropriate antimicrobial testing method, researchers can discern suitable conditions and streamline the identification of effective antimicrobial agents.
Introduction
Antimicrobial resistance (AMR) is a serious global concern, as infectious diseases become harder to treat [1]. Multi-resistant bacteria are on the rise, and the development of new antimicrobials is limited, making the situation even more challenging. To address this issue, researchers are exploring both natural and synthetic antimicrobial compounds as potential solutions [2]. Natural products, derived from plants, animals, and microorganisms, offer a rich source of biologically active compounds that show promise in combating infections. Additionally, synthetic antimicrobial compounds designed in laboratories have also demonstrated remarkable potential in fighting microbial infections. To unlock the potential of these compounds, appropriate antimicrobial screening and evaluation methods are essential. The in vitro antimicrobial assays play a crucial role in the discovery and development of new antimicrobial agents, providing crucial insights into their effectiveness and mechanisms of action. Understanding these methods better will help researchers identify potential antimicrobial candidates more efficiently and contribute to addressing the growing threat of AMR. While traditional technologies like disk diffusion and broth dilution methods are widely used, they may have limitations in reproducibility and time. On the other hand, newer methods like flow cytometry, bioluminescence, and impedance measurement offer higher sensitivity and throughput, but they may be costlier and less accessible in resource-limited regions. Thus, a comprehensive review of in vitro assays used to assess the antimicrobial activities of both natural and synthetic compounds is important.
This review provides an overview of the most common in vitro assays used to characterize the antimicrobial activity of promising natural and synthetic compounds. The methods covered include disk diffusion assay, well diffusion, spot assay, cross-streaking method, poisoned food method, agar dilution, broth macrodilution and microdilution, resazurin assay, co-culture method, time-kill kinetics, flow cytometry, thin layer chromatography-bioautography (TLC bioautography), bioluminescence assay and impedance measurement. For each assay, their respective advantages and limitations are presented, addressing issues of cost, accessibility, reproducibility, complexity of the tested samples, and other relevant factors. This comprehensive overview aims to aid researchers and practitioners in choosing appropriate screening techniques, allowing for the efficient identification of potential antimicrobial agents and contributing to the development of effective therapies against infectious diseases. By discussing gaps and limitations in current methodologies, the paper encourages researchers to explore approaches and technologies that enhance the accuracy, sensitivity, and efficiency of antimicrobial detection.
Sources of antimicrobials
The presence of antimicrobial activity has been documented in a diverse array of sources, encompassing natural origins derived from plants, animals, and microorganisms as well as synthetic compounds, nanoparticles, and so forth.
Plant sources of antimicrobials
Plants produce a diverse range of bioactive compounds as a part of their natural defense system. Many of these compounds, known as phytochemicals, are highly effective in combatting microbial pathogens. The phytochemicals employ various mechanisms to inhibit the growth and survival of microorganisms including bacteria, fungi, and viruses that make them promising candidates in developing antimicrobial agents. Among the phytochemicals, polyphenolics such as flavonoids, tannins, quinones, coumarins, and others have shown significant antimicrobial properties by disrupting microbial cell membranes, inhibiting key enzymes, and interfering with key cellular processes [3]. Terpenoids represent another important group of plant-extract bioactives which exhibit potent antimicrobial activity by disrupting microbial membranes, inhibiting protein synthesis, or interfering with essential metabolic pathways [4]. Some terpenoids such as essential oils have been found highly effective against a range of microorganisms rendering them valuable contenders for antimicrobial applications. Saponins, glycosides found in numerous plant species, also exhibit antimicrobial properties by disrupting microbial cell membranes [5]. Moreover, alkaloids, another group of plant-derived bioactive compounds, also demonstrated remarkable antimicrobial properties with some alkaloid-based drugs such as quinine, berberine, among others historically utilized for treating infectious diseases [3]. Additionally, plants also produce certain antimicrobial peptides (AMPs) such as defensins which also contribute to their antimicrobial activity by targeting microbial cell membranes [6]. Some plants produce lectins, which are carbohydrate-binding proteins with antimicrobial properties, offering another line of defense against pathogens. These diverse bioactive compounds derived from plant extract highlight their remarkable potential as sources of natural antimicrobial agents.
Animal sources of antimicrobials
The animal kingdom serve as a valuable resource for antimicrobial peptides and proteins which participate in their natural defense against harmful microorganisms. Like plants, various animal species including insects, amphibians, reptiles, birds, and mammals, also produce AMPs that can target and disrupt microorganisms' cell membranes, showing broad-spectrum activity against bacteria, fungi, and certain viruses [7]. For instance, amphibians like frogs and toads produce AMPs in their skin secretions, acting as a powerful defense against microbes. Defensins, the largest AMP family, are also produced by many animals, including mammals, birds, and insects. Defensins penetrate microbial membranes, leading to cell death and preventing infections in various anatomical sites like the respiratory and gastrointestinal tracts [8]. Additionally, animal sources provide other antimicrobial molecules like lysozymes, lactoferrin and related proteins [9]. Lysozymes, found in secretions like tears, saliva, and milk, break down bacterial cell walls causing bacterial cell lysis. Lactoferrin, an iron-binding protein in milk and bodily fluids, restricts microbial growth by sequestering essential nutrients like iron. Crustaceans and fish produce AMPs like piscidins and crustins which have demonstrated activity against bacteria, fungi, and viruses [10].
Microbial sources of antimicrobials
Microbial products encompass a remarkable diversity of antimicrobial activity, as microorganisms themselves produce potent compounds to compete for resources and ensure survival in their environments. Among them, antibiotics stand out as the most renowned and extensively studied natural antimicrobial compounds. Produced by certain bacteria and fungi, antibiotics possess the remarkable ability to inhibit the growth of other microorganisms. An iconic example is Penicillin, derived from the Penicillium fungus [11], which marked a groundbreaking advancement in modern medicine and paved the way for numerous crucial therapeutic agents to combat bacterial infections. Microorganisms also produce a large group of antimicrobial peptides known as bacteriocins. These small cationic peptides exhibit impressive effectiveness against closely related bacteria, contributing to food preservation by inhibiting spoilage and pathogenic bacteria in fermented foods [12, 13]. Additionally, the arsenal of microbial antimicrobial compounds includes other bioactive substances such as bacteriocin-like inhibitory substance (BLIS), cyclic lipopeptides, and lectins, all of which display potent activity against a wide range of microbial pathogens [14].
Some archaea and protists also produce antimicrobial compounds. For example, certain species of archaea produce antimicrobial peptides known as archaeocins which are active against other archaea and some bacteria, and are thought to play a role in inter-microbial competition [15]. Some species of protists, such as amoebae, also produce antimicrobial peptides and proteins that are active against bacteria [16].
Synthetic antimicrobials
Synthetic antimicrobial compounds are chemically synthesized substances meticulously designed to possess potent antimicrobial properties, providing protection against the escalating challenge of bacterial resistance to conventional antibiotics. Triclosan is an example of synthetic antimicrobial compounds that exhibit broad-spectrum antibacterial and antifungal activity [17]. Frequently used in personal care products like soaps and toothpaste, it disrupts bacterial cell membranes. Linezolid, another synthetic antibiotic, inhibits protein synthesis in Gram-positive bacteria including Staphylococcus aureus and Streptococcus pneumoniae [18]. Quinolones, a class of synthetic antimicrobial compounds such as ciprofloxacin and levofloxacin, impede bacterial DNA replication by inhibiting enzymes such as DNA gyrase and topoisomerase IV [19]. Additionally, synthetic peptides have been developed to mimic the natural antimicrobial peptides found in animals and plants. Pexiganan, one such synthetic peptide, exhibits antimicrobial activity against both Gram-positive and Gram-negative bacteria by disrupting bacterial cell membranes [20]. Other synthetic antimicrobial agents include daptomycin, a semisynthetic lipopeptide antibiotic that disrupts bacterial cell membranes and is used to treat skin and bloodstream infections caused by Gram-positive bacteria [21]; metronidazole, used to treat infections caused by anaerobic bacteria and parasites by disrupting their DNA synthesis; and others.
Antimicrobial nanoparticles
Nanoparticles, ranging from 1 to 100 nm in size, possess unique physicochemical properties due to their small size and large surface area. They can be produced synthetically, or naturally by microorganisms, plants, and animals. Nanoparticles exhibit potent antimicrobial activity against bacteria, fungi, and viruses. Their small size enhances the interaction with pathogens making them highly effective antimicrobial agents. Types of antimicrobial nanoparticles include silver nanoparticles which disrupt bacterial cell membranes and damage intracellular structures [22], zinc oxide nanoparticles which produce reactive oxygen species (ROS) elevating membrane lipid peroxidation that causes membrane leakage of reducing sugars, DNA, proteins, and reduces cell viability [23]; copper nanoparticles generate ROS, disrupt microbial cell walls and membranes, and interact with proteins and DNA [24]; the calcium oxide nanoparticles produce free radicals that damage bacterial cell membrane and the arrangement of polyunsaturated phospholipids [25]; and titanium dioxide nanoparticles which possess photocatalytic properties generating antimicrobial reactive oxygen species [26].
Methods to evaluate antimicrobial activity
The relentless quest for novel and highly effective antimicrobial compounds remains an ongoing pursuit driven by the pressing challenge of antimicrobial resistance, which diminishes the effectiveness of traditional antibiotics. To assess potential antimicrobial compounds, whether recently discovered or well-known, researchers employ a wide array of antimicrobial assay techniques. Among these, the following antimicrobial assay methods are extensively used for the identification and evaluation of antimicrobial activity.
1. Agar diffusion based screening of antimicrobial activity
Agar diffusion based assays, including the disk diffusion, well diffusion, agar plug, and agar spot assays, are widely utilized and cost-effective techniques in antimicrobial research for determining the antimicrobial activity of test compounds. These methods rely on the diffusion of antimicrobial agents from paper discs, wells, or plugs into the adjacent agar medium, inhibiting the growth of the test microorganism inoculated on the agar surface [27]. By measuring the resulting zone of inhibition (Fig. 1), which represents the area where microbial growth is prevented or inhibited by the agent, researchers can assess the relative potency of the test compound against the specific microorganism under investigation [28]. The agar diffusion assay serves as a valuable tool in antimicrobial research, providing qualitative data on the effectiveness of different substances against specific microorganisms.
Schematic representation of the disk diffusion and cross streak assay protocols in determination of antimicrobial activity. In the disk diffusion test, the indicator organism is evenly distributed on the agar surface (step 1). A disk containing an appropriate concentration of the antimicrobial agent is placed in the center of the plate (step 2). After incubation (step 3), the plate is examined for the presence of a clear zone of inhibition. In the cross-streak method, the test organism is initially streaked in a straight line on the agar surface (step 1). Following an incubation period (step 2), the indicator strains are streaked perpendicular to the test microbe (step 3). After another round of incubation (step 4), the plate is examined to observe the presence of clear zones at the intersections
Citation: European Journal of Microbiology and Immunology 14, 2; 10.1556/1886.2024.00035
The agar diffusion technique has a rich history dating back to the late 18th century, when Beijerinck first utilized it to study auxin’ effect on the growth of bacteria [29]. Since then, the method has undergone several modifications to enhance accuracy and efficiency, including the introduction of the ditch plate technique and the use of wells and filter paper discs. Furthermore, variations such as the agar plug and agar spot assays have been developed offering researchers flexibility and options. Although these variations employ the same principle of agar diffusion, they slightly differ in the method of introducing the antimicrobial agent into the agar medium.
Protocol
The protocol for the agar diffusion assay (Fig. 1) includes the following steps which typically takes 18–48 h to complete depending on the specific test microorganism [30]. Firstly, a culture of the test microorganism is prepared by inoculating it onto a suitable growth medium and incubating it under optimal conditions. Once the culture has reached the desired growth phase, a sterile cotton swab is used to evenly spread the microbial suspension across the surface of an agar plate, forming a uniform lawn. Next, wells of specific sizes are created in the agar using a sterile cork borer or a pipette tip and the test substance is carefully added to the respective well (well diffusion assay). Alternatively, instead of using wells, the agar spot assay offers the option to place filter paper disks (disk diffusion assay; also known as Kirby-Bauer method) or agar plugs (agar plug assay) impregnated with the test compound onto the agar surface. Another approach involves directly spotting a small volume of the test compound onto the agar surface using a sterile inoculating loop (agar spot assay). Subsequently, the plate is incubated at the appropriate temperature for a defined period to allow the compound to diffuse into the surrounding medium. Following incubation, the plates are carefully examined for the presence of inhibition zones. The area or diameter of the inhibition zones is measured to assess the antimicrobial activity of the tested substances [13]. Control plates containing only the solvent, culture filtrate or buffer are included to account for any potential effects of the vehicle on microbial growth.
Advantages
The agar diffusion assay has several advantages that make it a widely used method for screening antimicrobial activity [31]. It is a simple and cost-effective technique that requires minimal specialized equipment making it accessible to researchers with limited resources. The assay allows for the simultaneous evaluation of multiple test substances against a single microorganism, enabling comparative analysis of their antimicrobial activity. While the assay primarily provides qualitative information, the presence of clear zones surrounding the test substance can offer a relative indication of the potency of the antimicrobial agent. Moreover, the standardized protocol of the agar diffusion assay ensures relatively consistent results facilitating comparison of results across different studies and laboratories. In addition, this method is highly versatile as it can be used to test a wide range of antimicrobial agents including antibiotics, plant extracts, and synthetic compounds against diverse microorganisms. Furthermore, the assay provides a relatively rapid means of determining the susceptibility of microorganisms to a test compound with results typically available within 18–48 h. The agar diffusion assay also offers a few variations including the well, spot, plug, spot and the paper disc methods which provide additional flexibility and options for researchers. Lastly, the agar diffusion assay has been extensively validated and is recognized as a reliable method by regulatory authorities such as the Clinical and Laboratory Standards Institute (CLSI) [32].
Limitations
While the agar diffusion assay offers several advantages, researchers should be aware of some of its limitations which include its qualitative nature, variability in diffusion, restricted application, dependence on specific conditions, and subjectivity in interpretation [31, 33, 34]. The assay primarily produces qualitative results by indicating the presence or absence of antimicrobial activity through the formation of inhibition zones that serve as a relative measure of antimicrobial activity rather than providing precise quantitative measurements or minimum inhibitory concentrations (MICs). The size of the inhibition zone can be influenced by factors like temperature, humidity, and pH, affecting the diffusion rate of the test compound in the agar [1]. Additionally, variability in results can occur due to molecular weight, solubility, and diffusion rate, impacting the diffusion of test substances in the agar [31]. The assay can also be affected by differential diffusion of extract components in the aqueous media. This occurs when the antimicrobial extract, dissolved in a solvent, is impregnated into the paper disc or well on the agar plate. The chemical constituents in the extract may diffuse at different rates, resulting in uneven distribution of antimicrobial compounds around the disc or well. Moreover, the agar diffusion assay may have limitations in assessing the efficacy of certain antimicrobial agents, particularly those that are volatile or heat-labile. Furthermore, the assay's subjective interpretation of inhibition zones can introduce bias into the results, requiring training and experience for performing and interpreting the findings.
It is also important to note that the agar diffusion assay may not be as sensitive as other methods such as the microdilution assay in detecting low concentrations of antimicrobial agents. This means weak antimicrobial activity may not be detected, and the agar diffusion assay may require a sufficient quantity of the test substance which may not be practical for rare or expensive compounds. Researchers should also consider that the selection of specific microorganisms and the choice of growth media in the assay may not represent all relevant clinical or environmental conditions [33]. Despite these limitations, the agar diffusion assay remains a widely used and valuable technique for the preliminary screening of antimicrobial activity in test compounds.
2. Cross streak method
The cross-streak method is a unique variation of agar diffusion-based assays. In contrast to the previously described assays where the antimicrobial agent is placed on the agar medium, this method involves streaking the test microbe and the indicator microbe(s) across each other on the agar surface (Fig. 1). If the test microorganism produces an antimicrobial agent, it diffuses into the agar medium leading to inhibition of growth for the other microorganisms at the intersection points. Apart from its role in assessing antimicrobial activity, this method is also employed to study antagonism between microorganisms [35].
Protocol
The cross-streak method involves streaking of two different microorganisms across each other on an agar plate (Fig. 1) [36, 37]. Initially, a sterile inoculation loop or swab is used to streak the test microorganism in a straight line across the surface of an agar plate, typically positioned at the center. The plate is then incubated for a specific period based on the characteristics of the test microorganism, allowing for potential diffusion of any antimicrobial agents produced by the test microorganism into the agar medium. Following the incubation period, one or more microorganisms are streaked in separate straight lines that intersect the initial streak at 90-degree angle. The plate is once again incubated, and subsequent observations are made to evaluate the presence of growth-inhibition at the intersection of the two streaks. If the first microorganism produces an antimicrobial agent, it will diffuse into the surrounding agar medium, inhibiting the growth of the other microorganisms and resulting in the formation of clear zones of inhibition at the intersections.
Advantages
The cross-streak technique serves as a simple, fast, and cost-effective primary screening tool to assess antimicrobial activity. Its key advantage lies in its minimal requirement for specialized equipment and resources, rendering it an affordable and accessible method for antimicrobial assay. Moreover, the cross-streak method enables direct visual observation of the inhibition zone formed at the intersection of the streaks, facilitating a simple qualitative assessment of antimicrobial effectiveness. Additionally, the technique allows for the concurrent evaluation of a test microorganism's capability to inhibit the growth of multiple microorganisms on a single agar plate. Furthermore, by streaking different microbes perpendicular to the initial streak of the test microorganism, it enables comparative assessments in a single experiment [38].
Limitations
The cross-streak method, like the agar diffusion assays, is subject to similar limitations related to the diffusion of the antimicrobial compounds through the agar. These limitations include variations in the size of the inhibition zone due to factors such as the diffusion properties of the agent, concentration used, and susceptibility of the target microorganisms. Additionally, the method primarily provides qualitative data and does not yield quantitative information on the potency of the antimicrobial agent [39]. The subjective nature of the method, as the interpretation of the inhibition zone relies on visual observation, may introduce potential bias or variability when determining the presence or absence of growth inhibition.
3. Co-culture assay
The co-culture assay involves simultaneous growth of two microbes in a shared environment facilitating direct interaction between them like the cross-streak method. Growth of the indicator organism is subsequently assessed and compared to a control to confirm the inhibitory activity of the test microbe [40]. This method is commonly employed to assess the antimicrobial effects of probiotic lactic acid bacteria against pathogens.
Protocol
In the co-culture assay, the microbe of interest and an indicator strain are cultured together in a growth medium that supports the growth of both bacteria. For instance, when assessing the antimicrobial effect of a lactic acid bacterium against Escherichia coli, a modified MRS media (mMRS) consisting of MRS broth and nutrient broth is typically employed to facilitate the growth of the lactic acid bacteria and E. coli, respectively [41]. A positive control with a known microbe exhibiting antimicrobial activity and a negative control with a microbe lacking antimicrobial activity are included. At specific intervals, an aliquot from the broth is taken out and spread on selective agar media that exclusively supports the growth of the indicator strain. After incubation, colony counting is performed to determine the viable count. If the growth of the test microbe is lower than that of the negative control or if its growth does not increase over time, it indicates the production and secretion of antimicrobial compounds by the test microbe.
Advantages
One of the key strengths of the co-culture assay is its potential to replicate microbial interactions, providing a more realistic assessment of the antimicrobial activity of test microbes compared to some other methods. While it may not fully mimic the complexities of the natural environment, it enables direct observation of the interaction between the test and indicator organisms in the same broth in real time. It also supports the evaluation of microbial antagonism and has the potential to differentiate between bactericidal and bacteriostatic effects based on the growth patterns compared to the positive and negative controls. The assay can also provide valuable information on the potential synergistic or antagonistic interactions between the microorganisms. Additionally, by assessing bacterial growth and viability at multiple time points, this method allows for the observation of time-dependent changes in antimicrobial activity. The assay is relatively simple and cost-effective, requiring minimal specialized equipment or expertise.
Limitations
The co-culture assay is not suitable for testing potency of the antimicrobial activity. It is designed to screen the antimicrobial activity of one microbe against another, making it less suitable for evaluating other types of antimicrobial sources like plant extracts which are rather assessed using variants of this test such as the time kill assay. The use of two different media in a single broth-culture can potentially interfere with the growth of both microbes and may not fully reflect the antimicrobial activity, potentially impacting the reliability and reproducibility of results. Additionally, microbial interactions may involve the exchange of metabolites or nutrients leading to cross-feeding effects that could potentially make it more challenging to interpret antimicrobial activity in co-culture assays [42].
4. Poisoned food technique
The poisoned food technique is an antimicrobial screening method that has been utilized for centuries, primarily focusing on evaluating antifungal activity [43]. In contrast to agar diffusion assays, where the test substance diffuses from a paper disk into agar, the poisoned food method involves direct mixing of the test substance with the growth medium where the test microorganism is subsequently placed. The growth inhibition of the test organism is then assessed. This technique has been widely employed in the investigation of antimicrobial activity in plant extracts. While the poisoned food technique is still utilized today, it is gradually being supplanted by more sensitive methods. Nonetheless, it remains a valuable tool for screening antimicrobial activity, especially within the context of evaluating natural products, extracts, and synthetic compounds against microorganisms.
Protocol
To perform the assay, agar plates containing desired amounts of the antimicrobial compound of interest and control plates devoid of the test compound are prepared [44]. Agar discs (3–6 mm) containing the test fungus are cut from the periphery of actively growing colonies and carefully placed at the center of the petri plates. The plates are then incubated at an appropriate temperature for a specific duration. After incubation, the growth of the microorganisms on both plates is compared, and if there is significantly less microbial growth on the treated plate compared to the control, it indicates that the substance being tested is effective in inhibiting the growth of microorganisms. The antifungal effect (%) is expressed as a percentage, indicating the degree of inhibition in fungal growth caused by the test substance calculated using the following equation: (growth diameter in control – growth diameter in test)/growth diameter in control × 100 [45].
Advantages
The poisoned food technique is a cost-effective and simple method that is particularly useful for testing the antifungal activity of substances. It offers a straightforward approach that can be performed with ease, requiring minimal specialized equipment or materials.
Limitations
As a preliminary screening method, the poisoned food technique may not provide precise quantitative data regarding the potency or MIC of the antimicrobial agent. It is also important to note that this method is primarily employed for evaluating antifungal activity and is less commonly utilized for screening antibacterial activity.
5. Time kill kinetics
The time kill kinetics assay evaluates antimicrobial activity by exposing the test microorganism to different concentrations of the antimicrobial agent over a specific duration. It provides valuable information about the time-dependent or concentration-dependent effects of the agent and offers information on the temporal dynamics of antimicrobial activity. The time kill assays are employed to assess novel antimicrobial agents and combinations of these agents [46].
Protocol
To assess time kill kinetics, a series of test tubes or wells containing a growth medium are prepared, with each tube or well containing a specific concentration of the antimicrobial agent. The test microorganism is then inoculated into each tube or well, ensuring a consistent starting cell density. The tubes or wells are incubated under controlled conditions for predetermined time intervals. At each interval, samples are withdrawn from each tube or well and plated onto appropriate growth media. After incubation, the number of viable microorganisms is determined by counting the colonies. The results obtained from each time point are analyzed to assess the impact of the antimicrobial agent on the growth and viability of the microorganism over the specified time period.
Advantages
The time kill kinetics assay is a simple but informative approach to assess the efficacy of antimicrobial agents, aiding in the determination of optimal dosing regimens and exposure durations. Unlike static assays that provide a single time point measurement, the time kill kinetics assay allows for the continuous monitoring of bacterial growth providing information on the rate and extent of microbial killing or growth inhibition. By measuring the bacterial population at different time intervals, this assay offers valuable insights into the antimicrobial efficacy, kinetics, and potential bactericidal or bacteriostatic effects of the tested substance [47]. Time-kill kinetics can be used to evaluate the combined effects of multiple antimicrobial agents on microbial growth. This can help to identify synergistic or antagonistic interactions among antimicrobial compounds potentially enhancing the efficacy of antimicrobial therapy.
Limitations
The time kill kinetics assay, while valuable, has certain considerations. It may be time-consuming and resource-intensive due to the requirement for continuous monitoring and sampling. The assay may not provide extensive insights into specific microbial killing mechanisms. Moreover, as an in vitro approach, it may not fully replicate the intricate microbial interactions present in the host environment. The assay evaluates antimicrobial agents at static concentrations, which may not precisely mimic the dynamic drug concentrations observed in vivo [48].
6. Agar dilution and broth dilution methods
The agar dilution and broth dilution methods are two highly similar techniques that are used for determining the MIC of antimicrobial agents. The MIC is the lowest concentration of an antimicrobial agent that inhibits the visible growth of a particular bacterium (Fig. 2). The basic difference between the agar dilution and broth dilution methods lies in the medium used: agar dilution employs agar plates, whereas broth dilution utilizes liquid broth tubes. Both methods involves incorporating different concentrations of the antimicrobial substance into the medium, followed by the application of a standardized number of microbial cells to the medium. The MIC is determined by observing the growth or absence of growth of the microorganism. The agar dilution and broth dilution methods are both important for guiding the selection of appropriate antimicrobial therapy and monitoring changes in antimicrobial susceptibility over time.
Agar dilution and broth macrodilution protocols for antimicrobial evaluation. In agar dilution, the indicator microbe is inoculated on agar plates having varying concentrations of the antimicrobial agent. The minimum inhibitory concentration (MIC) is determined as the lowest concentration of the agent preventing visible growth after incubation. In broth macrodilution, the indicator microbe is inoculated into broth tubes containing different concentrations of the antimicrobial agent. The MIC is determined as the lowest concentration with no visible growth
Citation: European Journal of Microbiology and Immunology 14, 2; 10.1556/1886.2024.00035
Protocol
In the agar dilution method, a series of agar plates are prepared containing a concentration gradient of the antimicrobial agent being tested (Fig. 2). A standardized inoculum of one or more test microorganisms is then inoculated onto the agar surface of each plate. Following appropriate incubation, the plates are carefully examined for the presence of microbial growth. The MIC is determined by identifying the lowest concentration of the antimicrobial agent that completely inhibits visible growth of the microorganism on the agar surface.
In contrast, the broth dilution method has two variants based on the volume of the reaction mixture: macrodilution and microdilution. Broth macrodilution is conducted in a series of tubes with a larger volume, while broth microdilution is performed in wells of a microtiter plate with a smaller volume. The tubes or wells are filled with liquid media containing different concentrations of the antimicrobial agent (Fig. 2). A standardized inoculum of the test microorganism is then added to each tube or well. Subsequently, the tubes or microtiter plate is incubated under controlled conditions and the growth of the test microbe is assessed using a spectrophotometer or microplate reader. The MIC is determined as the lowest concentration of the antimicrobial agent that visibly inhibits the growth of the microorganism in the broth.
Advantages
The agar and broth dilution methods have been standardized by international organizations such as the Clinical and Laboratory Standards Institute (CLSI) and the European Committee on Antimicrobial Susceptibility Testing (EUCAST), and are widely accepted as reliable methods for determining antimicrobial susceptibility. The agar dilution method is considered the gold standard for antimicrobial susceptibility testing due to its accuracy and consistency. It is often used to test the effectiveness of new antibiotics against a large panel of bacteria. One advantage of the agar dilution method is its ability to visually check for bacterial growth on agar plates, allowing for convenient detection without relying on changes in opacity or color [49]. This simplifies the determination of bacterial growth compared to assessing turbidity changes in the broth dilution technique. When the medium is strongly colored or opaque, it provides a simpler and clearer means to identify bacterial growth on the agar surface.
Moreover, the agar dilution method can be enhanced in efficiency by incorporating semi-automation through the use of an inoculum-replicating apparatus. This technological advancement facilitates the consistent and precise transfer of bacterial inocula to multiple agar plates, reducing manual labor and potential human errors. Additionally, the agar dilution method offers the advantage of testing multiple bacteria simultaneously on the same sets of plates, whereas the broth dilution method enables testing multiple antimicrobial agents at a time against a single microbe [50]. Both agar and broth dilution methods overcome the limitations of the disk diffusion method by providing quantitative data through MIC determination, facilitating more meaningful conclusions [51]. These methods also have the potential to improve the identification of MIC endpoints and extend the range of antibiotic concentrations. Additionally, the broth microdilution method performed in the microtiter plate format offers more flexibility, speed, cost minimization, and convenience [52].
Limitations
The agar and broth dilution tests can be time-consuming because they require the preparation of a series of agar plates or broth tubes each having a different concentration of the antimicrobial agent [53]. Manual preparation of the serial dilutions can also be tedious, particularly if not automated. Additionally, agar plates have a short shelf-life and must be used within a week of preparation. Moreover, the agar dilution method can test only one antimicrobial agent at a time, while the broth dilution method can assess only one indicator organism at a time. Furthermore, these methods may not be suitable for testing the susceptibility of fastidious or slow-growing bacterial species, and may not be appropriate for all types of sample types [54]. For instance, in agar dilution, hydrophobic extracts may separate out from the agar, while in broth dilution, essential oils may not remain in solution for the duration of the assay, and emulsifiers and solvents may interfere with the accuracy of results. It is important to note that the MIC values obtained from these methods do not indicate the mode of action of the antimicrobial agent and may not accurately predict the drug's efficacy in vivo [55]. While these limitations exist, these methods remain valuable tools in antimicrobial evaluation and are often recommended as a standardized antimicrobial susceptibility testing method [56].
7. Resazurin assay
The resazurin assay is a versatile colorimetric test that measures the metabolic activity of living cells. The principle of the test is based on the reduction of resazurin, a blue-colored non-fluorescent dye, to resorufin, a pink-colored fluorescent dye, by the metabolic activity of living cells (Fig. 3). The amount of resorufin produced is directly proportional to the metabolic activity of the cells, and can be quantified by measuring the fluorescence or absorbance of the solution. The application of the resazurin test spans various fields, including microbiology, cell biology, and drug discovery. In microbiology, the test is commonly used to determine bacterial growth and antimicrobial susceptibility, and is often used in conjunction with the microdilution assay. This allows for the rapid and quantitative assessment of bacterial viability, and is a valuable tool for evaluating the efficacy of antimicrobial agents. Additionally, the resazurin assay has been increasingly used for determining the antimicrobial activity of nanomaterials and plant extracts [57–59].
Resazurin assay protocol for screening and evaluation of antimicrobial activity. In the resazurin test, the antimicrobial agent is added to the wells of a microtiter plate. The plate is then inoculated with the target microorganism and incubated. Resazurin is subsequently introduced to the wells and incubated further. The color change, from blue to pink, indicates the presence of viable cells. The minimum inhibitory concentration (MIC) can be determined as the lowest concentration of the antimicrobial agent that prevents the observed color change
Citation: European Journal of Microbiology and Immunology 14, 2; 10.1556/1886.2024.00035
Protocol
The resazurin assay protocol involves dispensing the desired concentrations of the antimicrobial agent into the wells of a 96-well microtiter plate (Fig. 3) [58]. A standardized cell suspension of the target microorganism, prepared in a suitable broth, is then added to the wells. The microplate is subsequently incubated at the specific temperature and duration appropriate for the microorganism being tested, allowing for its growth in the presence of the antimicrobial compounds. Subsequently, the resazurin solution is introduced to each well, ensuring an optimized final concentration based on the experimental conditions. The mixture is further incubated for 3–4 h. Following this incubation period, the wells are examined for any color changes. A shift from blue to pink or purple indicates the presence of viable cells, signifying metabolic activity and cell viability. Conversely, a lack of color change or minimal alteration suggests inhibition or killing of the microorganisms. To quantify the results, the absorbance or fluorescence of the wells is measured using a microplate reader. The intensity of the signal reflects the metabolic activity and viability of the microorganisms. The assay also allows for the determination of MIC by ascertaining the lowest concentration of the antimicrobial agent that effectively prevents the observed color change [60].
Advantages
The resazurin assay is a versatile and reliable method for evaluating antimicrobial properties. It offers several advantages including simplicity, sensitivity, rapidity, and robustness [58]. This assay allows for the detection of microbial growth in small volumes of solution within microtiter plates. Compared to the OD method, it exhibits higher sensitivity and offers early detection of microbial inhibition [61]. By monitoring metabolic activity and microbial viability, this test produces a distinct color change from blue to pink providing a clear indication of the presence or absence of viable microorganisms. The method is versatile and can be utilized to evaluate the antimicrobial activity of diverse antimicrobial agents against a wide range of microorganisms including both bacteria and fungi. It is a cost-effective technique requiring minimal equipment, and its high-throughput setup allows for simultaneous screening and assessment of multiple antimicrobial agent-microbe combinations. The assay also enables the quantification of antimicrobial activity through the determination of the MIC. Moreover, the non-invasive nature of the resazurin assay helps preserve the integrity of the compounds of interest facilitating further analysis.
Limitations
Resazurin reduction is strongly linked to oxygen consumption, making it a preferred assay for aerobic or microaerophilic microorganisms, while strictly anaerobic microorganisms are less likely to reduce it [62]. Another limitation is its toxicity towards certain cells [63, 64], although the concentration used in the assay is typically lower than the levels inhibitory to bacteria. Care should be taken when using resazurin in the presence of nanomaterials, as they can influence its reduction and fluorescence. Additionally, high bacterial concentrations or long incubation periods can lead to extensive reduction of resazurin which can interfere with accurate measurement of the antimicrobial activity [61]. This can be overcome by optimizing the inoculum size and incubation time. Another study found that the color change in the resazurin assay took longer in colistin-resistant Pseudomonas aeruginosa compared to other bacteria [65], suggesting that the observation time needs to be adjusted based on the target strain.
8. Antimicrobial gradient diffusion test
The antimicrobial gradient diffusion test offers an alternative approach for evaluating antimicrobial activity by combining the fundamental principles of dilution and diffusion of the antimicrobial agent [66]. Unlike agar diffusion assays, which primarily serve as screening tools, the gradient diffusion test is specifically designed to determine the MIC. This method involves the dilution of the antimicrobial agent on a specialized test strip (Fig. 4), thereby creating a concentration gradient that spans a range of concentrations. Following inoculation of agar plates with the test organism, the test strips are positioned atop the agar surface. Subsequently, the antimicrobial agent gradually diffuses from the strip into the agar, leading to the formation of a discernible zone of inhibition. Developed in the 1980s, this method has gained popularity as an alternative to the agar dilution method for assessing the MIC of antibiotics against bacteria. It is worth noting that the test strips employed in the gradient diffusion test are predominantly commercially available, widely known as E-test strips. These strips are primarily employed for the precise determination of antibiotic susceptibility among pathogens, rather than serving as tools for general screening of antimicrobial activity or evaluating the efficacy of novel antimicrobial agents. The E-test method assumes a pivotal role in clinical microbiology laboratories, effectively guiding treatment decisions and facilitating the judicious selection of appropriate antibiotics to combat bacterial infections [67].
The gradient diffusion E-test method of antimicrobial assay. An E-test strip with a predefined gradient of antimicrobial agent is placed on an agar plate inoculated with the test microorganism. As the antimicrobial agent diffuses into the agar, an ellipse zone of inhibition is formed. The minimum inhibitory concentration (MIC) is determined by visually identifying the point where the zone intersects the scale on the strip
Citation: European Journal of Microbiology and Immunology 14, 2; 10.1556/1886.2024.00035
Protocol
To perform the gradient diffusion test, the agar media is uniformly inoculated with the test organism using a sterile swab, ensuring even and consistent distribution of the inoculum. Next, a test strip, which consists of a plastic strip impregnated with a predefined gradient of the antimicrobial agent, is carefully placed on the agar surface [68]. The strip is pressed lightly to ensure contact with the agar. It is possible to accommodate multiple test strips on the agar plate. For example, up to five or six test strips can be arranged in a radial pattern on the surface of a 150-mm agar plate. During incubation, the antimicrobial agent diffuses from the strip into the agar, creating a concentration gradient. Over time, the antimicrobial agent inhibits the growth of the organism, resulting in a characteristic elliptical zone of inhibition (Fig. 4). After the appropriate incubation period, typically 16–18 h, the plate is examined. The concentration of the antimicrobial agent can be directly read from the scale printed on the upper side of the E-test strip. The MIC is determined by identifying the point where the zone of inhibition intersects the E-test strip.
Advantages
The gradient diffusion test is widely acknowledged as a convenient and rapid approach for conducting antimicrobial susceptibility testing [51]. It exhibits excellent reproducibility, ensuring consistent and reliable results. Moreover, multiple antimicrobial agent can be tested on the same agar plate at a time. The test provides a quantitative assessment of the MIC, allowing for a precise evaluation of the antimicrobial agent. The predefined gradient on the E-test strip also facilitates visualization and easy interpretation of the MIC value. Moreover, studies have shown a significant correlation between E-test results and MIC values obtained through broth or agar dilution methods [69, 70].
One key advantage of the E-test strip is the stability of the concentration gradient which enables its use in evaluating antimicrobial susceptibility of various types of bacteria including fastidious and slow-growing organisms, as well as a diverse range of clinically significant organisms across multiple species [71]. Additionally, the E-test can also be utilized to assess the susceptibility of yeast, molds, and mycobacteria [72, 73]. In the disk diffusion assay, certain antibiotics pose challenges due to their specific physicochemical properties [73]. For example, glycopeptide antibiotics, like vancomycin, have a large molecular weight which hinders their diffusion in agar. This limited diffusion and poorly defined concentration gradient around a vancomycin disk can lead to only a slight difference in zone size between susceptible and resistant strains, potentially resulting in unreliable results. To overcome this limitation, the Centers for Disease Control and Prevention (CDC) recommend using an MIC method, such as the E-test, for vancomycin susceptibility testing in staphylococci. The E-test enables the determination of the MIC of vancomycin, which helps to detect reduced susceptibility to the antibiotic, even in strains that may not show a significant difference in zone size in the disk diffusion method. While standardized MIC methods are effective in detecting strains with higher levels of resistance to vancomycin (MIC ≤8 μg mL−1), strains with lower MIC values (4–8 μg mL−1), known as intermediate and/or heterogeneously resistant strains, can be better detected using a more nutrient-rich medium like brain-heart infusion agar, a higher inoculum, and extended incubation, conditions described in the E-test macro-method for the detection of vancomycin intermediate Staphylococcus aureus (VISA) and hetero-VISA (h-VISA) [73].
Limitations
The gradient strip diffusion test is primarily designed for testing susceptibility to antibiotics and may not be suitable for other types of antimicrobial agents or novel compounds. The cost of E-test strips can be higher compared to some other susceptibility testing methods, making it less economically feasible for large-scale testing or resource-limited settings [74]. Additionally, the interpretation of the gradient strip test results may be subjective, as it relies on visual inspection to determine the MIC value. This subjectivity could introduce variability in the interpretation of results among different observers. Moreover, interpreting the endpoints in the strip diffusion test can be challenging due to the possibility of feathery or swarming edges produced by certain strains at the intersection with the test strip. The E-test method has also been observed to exhibit systematic biases towards higher or lower MIC values for certain microorganism-antimicrobial agent combinations [74, 75]. The method also demonstrated inaccurate and inconsistent behavior when assessing certain antibacterial agents, including penicillin, ciprofloxacin, ofloxacin, and rifampicin [76].
9. TLC-bioautography
TLC-bioautography is a highly effective technique widely employed in natural product research for the screening and assessment of bioactive compounds including antimicrobial agents [77, 78]. The fundamental principle of TLC-bioautography is based on the separation of components in a mixture using a TLC plate, followed by a direct bioassay performed on the plate for component detection (Fig. 5). This technique allows for the rapid and efficient identification of bioactive compounds in complex mixtures. TLC-bioautography has found extensive application in evaluating the antimicrobial activity of diverse natural products, encompassing essential oils, plant extracts, and bacterial secondary metabolites.
Different formats of the thin-layer chromatography (TLC)-bioautography assay utilized for antimicrobial activity determination. In the direct bioautography variant, a microbial broth is sprayed or dipped onto the developed TLC plate. For agar diffusion or contact bioautography, the developed plate is placed onto a petri dish with agar pre-seeded with the target microorganism, and after 30–60 min, the TLC plate is removed, while the agar plate continues incubation. In immersion or overlay bioautography, the TLC plate is immersed or overlaid with a seeded molten agar medium. All three methods enable the detection of active compounds as zones of inhibition
Citation: European Journal of Microbiology and Immunology 14, 2; 10.1556/1886.2024.00035
Protocol
There are three main variants of TLC-bioautography, each differing in the method by which bacteria are introduced to the TLC plate: agar diffusion or contact bioautography, immersion or overlay bioautography, and direct TLC-bioautography (Fig. 5) [78]. Each variant follows the initial step of conducting TLC to separate the components of the sample mixture. In TLC, a small amount of the sample is spotted on a TLC plate, which is then developed in a solvent system. The components in the sample mixture separate based on their physical and chemical properties, resulting in distinct spots on the TLC plate which can be visualized using UV light or by staining the plate with a suitable reagent. Once the TLC plate has been developed, one of the three variants of bioautography can be performed to identify compounds with antimicrobial activity.
In the direct bioautography variant, the developed TLC plate is directly sprayed with or dipped into a microbial broth (Fig. 5). Following incubation, the active compounds are detected as zones of inhibition or growth inhibition on the TLC plate. This approach is currently the most widely utilized among the three variants.
In agar diffusion or contact bioautography, the developed plate is placed onto a petri dish containing a layer of agar pre-seeded with the target microorganism (Fig. 5). The TLC plate is left undisturbed for approximately 30 min to an hour, enabling the diffusion of compounds into the agar. Subsequently, the TLC plate is removed, and the agar medium is incubated under appropriate conditions. The compounds from the TLC plate diffuse into the agar medium and interact with the microorganism. The presence of antimicrobial compounds inhibits the growth of the indicator organism, resulting in the formation of distinct clear zones (bioautograms) on the agar surface. Following incubation, the plate is examined for growth inhibition zones, which indicate the interaction between the compounds and the microorganism. These zones correspond to the location of bioactive compounds on the TLC plate.
In immersion or overlay bioautography, the developed TLC plate is immersed or overlaid with a seeded molten agar medium (Fig. 5). After solidification of the medium and incubation, the bioactive compounds migrate from the TLC plate into the agar medium, inhibiting the growth of the indicator organism and forming bioautograms which indicate the presence and location of bioactive compounds on the TLC plate. This method serves as a valuable alternative when direct bioautography is unfeasible, particularly in cases where mycelia blockage obstructs the spray head or when it is necessary to minimize microbial dissemination and contamination within the laboratory. Nonetheless, it is important to note that the sensitivity of this method is relatively limited due to the dispersion and dilution of active substances that occur throughout the agar gel matrix.
The inhibition zones in each method are naturally observed for spore-producing fungi and pigmented bacteria or can be revealed through the use of tetrazolium salts. The compounds can be identified by comparing their retention factor (Rf) values on the TLC plate to the Rf values of known standards.
Advantages
TLC-bioautography is a powerful technique for the identification of antimicrobial compounds due to its capability to precisely localize activity within complex matrices [79]. This technique enables simultaneous detection and targeted isolation of the bioactive constituents. Bioautography provides a direct visualization of bioactive compounds by incorporating the biological component into the analytical procedure. This allows for the identification and localization of unknown antimicrobial compounds within a sample mixture [77]. The assay is relatively simple and yet sensitive, effectively detecting antimicrobial compounds even at low concentrations [80]. It offers a rapid screening method, enabling the detection of bioactive components in a short period of time. Moreover, bioautography can be applied to various sample types including plant extracts, natural products, and synthetic compounds, making it a versatile tool for antimicrobial activity assessment.
Limitations
One limitation of bioautography is its qualitative nature, lacking the capacity to provide quantitative data on compound activity [81]. Consequently, it can be challenging to compare the activity of different compounds or to determine the optimal concentration of a compound for its activity. Additionally, the sensitivity and reproducibility of results depend on the growth conditions of the microorganisms used in the assay. Some antimicrobial compounds in a sample may remain undetected if they are not active under the assay's conditions. Interference from other compounds within the sample matrix can also affect the bioautography assay, making it difficult to identify active compounds accurately. Furthermore, bioautography is limited by its dependence on the extraction method and the choice of the TLC solvent [54].
10. Flow cytometry
The basic principle of flow cytometry involves the use of lasers to detect and measure the physical and chemical characteristics of cells or particles in a fluid stream. The technique allows for the rapid and simultaneous analysis of multiple parameters, including cell size, granularity, and fluorescence. In the context of antimicrobial activity, flow cytometry can be used to assess the effects of antimicrobial agents on microbial cells, including changes in cell viability, membrane integrity, and metabolic activity [82]. For example, flow cytometry can be used to measure the percentage of live and dead cells in a bacterial culture following treatment with an antimicrobial agent, providing a rapid and accurate assessment of the agent's activity.
Protocol
A typical protocol may involve exposing the target microbe to the antimicrobial agent for a specified period of time, then staining the cells including control samples with specific fluorescent dyes targeting viability markers or cellular functions [83]. Common dyes like SYTO 9 and propidium iodide (PI) can distinguish between live and dead cells based on their membrane integrity [84]. The stained cell suspension is then passed through a flow cytometer instrument, where it is directed in a single-file manner through a laser beam. As the cells pass through the laser, they scatter light and emit fluorescence signals, which are captured by detectors. Analyzing this signal enables the determination of cell viability, membrane integrity, metabolic activity, and other relevant characteristics. The instrument yields quantitative data on the percentage of viable and non-viable cells, as well as the overall dynamics of the microbial population. The obtained data can be further analyzed to evaluate the antimicrobial effects of the tested agents. By comparing treated and untreated samples, researchers can assess the impact of antimicrobial agents on microbial populations.
Advantages
Flow cytometry provides unique advantages for studying antimicrobial activities, allowing researchers to assess the effects of selected compounds on thousands of individual cells within seconds [85]. Its ability to conduct rapid and high-throughput analysis of large cell populations facilitates efficient screening of potential antimicrobial agents. The technique also offers high precision, accuracy and sensitivity [83] allowing for the detection of subtle changes in cell viability and providing valuable information on the efficacy of antimicrobial agents. Moreover, flow cytometry has the capability to discriminate between live and dead cells based on their membrane integrity, which further enhances our understanding of the mechanisms of antimicrobial agents [86]. This technique is versatile and applicable to various types of microorganisms including bacteria, fungi, and parasites, making it a valuable tool in antimicrobial research and drug development. The ability to assess antimicrobial activity at the single-cell level and obtain quantitative data also enhances the reliability of the results.
Limitations
Flow cytometry necessitates specialized equipment and expertise, which can incur considerable costs and may not be readily accessible in all research environments. Substantial initial investment and extensive training are required for equipment calibration, complex assay setup, and data analysis [87]. The expenditure on reagents and maintenance can also be substantial. The intricate nature of flow cytometry analysis may present a steep learning curve for researchers, leading to potential errors or data misinterpretation. Additionally, the technique's sensitivity to various experimental variables, such as sample preparation and instrument settings, may impact result reproducibility. The use of fluorescent labels or specific probes to assess certain microbial properties can influence cell behavior and introduce artifacts, potentially affecting antimicrobial activity measurements. Furthermore, while flow cytometry is well-suited for evaluating planktonic microbial cells, its application to study antimicrobial activity against biofilms may necessitate additional considerations and adaptations [88].
11. Bioluminiscence assay
The principle of the bioluminescence assay involves the use of genetically engineered bacteria that express a light-emitting enzyme, commonly luciferase [89]. These bacteria are designed to produce light in response to a specific metabolic process, such as ATP production. ATP, being present in all living and metabolically active cells, serves as a reliable indicator of viable microbial cells. The luciferase enzyme catalyzes the luciferin-luciferase reaction, where luciferin, a light-emitting molecule, undergoes oxidation in the presence of oxygen, ATP, and magnesium ions, leading to the production of oxyluciferin and light. When the bacteria are exposed to an antimicrobial agent, the metabolic processes are disrupted, resulting in a decrease in light production. The amount of light produced over time can be measured and used to determine the effectiveness of the antimicrobial agent against microbes. The luciferase reaction plays a pivotal role in this assay, enabling the detection and quantification of ATP levels in live microbial cells, providing valuable information about their metabolic activity and viability in response to antimicrobial agents.
Protocol
In bioluminiscence assay, the target microorganism is genetically engineered to express bioluminescent proteins, typically luciferase, by introducing specific genes or plasmids into their genomes [89]. Subsequently, the bioluminescent strain is cultured and treated with the antimicrobial agent under investigation. The treated microbial cells are incubated, and bioluminescent signals are detected using a luminometer or a bioluminescence imaging system which allows for the measurement of the emitted bioluminescent light over a defined period of time. The emitted light intensity is proportional to the ATP levels in the cells, reflecting their metabolic activity and viability. The collected bioluminescence data is analyzed to determine the effects of the antimicrobial agents on the microbial population. The reduction in bioluminescent signals indicates the inhibition of microbial growth and the effectiveness of the antimicrobial agents.
Advantages
The bioluminescence assay is highly versatile and can be adapted to a wide range of microorganisms and antimicrobial agents. It provides high sensitivity, allowing for the detection of even subtle changes in microbial viability and metabolic activity. The assay provides real-time, non-invasive and continuous monitoring of antimicrobial effects [90, 91], enabling researchers to observe dynamic changes in microbial populations during treatment. Furthermore, its quantitative nature allows for precise measurements of antimicrobial efficacy.
Limitations
The bioluminescence assay requires for genetic engineering or transformation of the microorganisms with the luciferase gene. This process can be time-consuming and labor-intensive, making it relatively less practical for routine antimicrobial screening, especially when dealing with a large number of different microorganisms. Moreover, the interpretation of bioluminescence assay results can be complex, as the relationship between light production and bacterial growth or viability may not always be straightforward. Another drawback is the quenching of emitted light, which can negatively impact the measurements. Specifically, certain compounds present in the biological samples can significantly diminish the photometrically measured light. Conversely, some non-microbial substances in the biological samples that emit light can enhance the intensity of the measured light [92]. Luciferase markers may also have certain limitations, such as their reliance on the concentration of available substrate, the physiological condition of the bacteria, the limited stability of the luciferase enzyme, and the existence of factors that can either stimulate or inhibit the sample being measured, producing false readouts [93, 94].
12. Impedance measurement
The impedance-based assay is a powerful technique that can detect alterations in the electrical properties of microbial cells in response to interactions with antimicrobial agents [95]. By measuring changes in electrical impedance, researchers can infer valuable information about cell volume, membrane permeability, and metabolic activity [96]. These recorded changes are then carefully analyzed to determine the effectiveness of the antimicrobial agent being tested. The real-time and label-free nature of impedance measurement makes it a valuable tool in antimicrobial screening, offering insights into the dynamic response of microbial cells to potential treatments.
Protocol
The basic protocol involves placing the microbial sample in a specialized chamber with electrodes to which an alternating current is applied [95, 96]. As the current passes through the sample, impedance changes are recorded at different frequencies. The impedance values are then analyzed to determine the effects of the antimicrobial agents on the microbial cells, such as changes in cell viability, growth, and metabolism. This real-time and label-free approach allows researchers to rapidly assess the antimicrobial activity of test compounds and provides valuable data for screening and evaluating potential antimicrobial agents.
Advantages
The impedance-based antimicrobial assessment offers real-time and label-free monitoring of microbial responses to antimicrobial agents, enabling researchers to observe changes in cell properties without the need for additional labels or stains [97]. This method exhibits high sensitivity in detecting microbial growth and the effects of antimicrobial agents by identifying even subtle changes in impedance. The rapidity of impedance measurements allows for high-throughput screening of multiple compounds in a short time [95], making it an efficient approach in the early stages of antimicrobial drug discovery.
Limitations
Impedance measurement requires specialized equipment and setup which may not be readily available in all research settings, leading to their higher costs and limited accessibility. The interpretation of impedance data can be challenging, and the lack of standardized protocols may result in variations in experimental procedures and data analysis [97]. The validation and standardization of impedance-based methods for antimicrobial screening and evaluation are still ongoing, and further research is needed to establish their reliability and reproducibility for routine analysis.
Other emerging techniques
In addition to the methods mentioned earlier, emerging techniques such as mass spectrometry, and time-lapse microscopy show great promise for the characterization and analysis of antimicrobial compounds, although they are less commonly employed for high-throughput screening and large-scale evaluation of antimicrobial activity. Mass spectrometry enables real-time monitoring of volatile organic compounds (VOCs) produced during microbial metabolism, offering dynamic insights into antimicrobial effects. By analyzing the diverse VOC profiles, researchers can gain a deeper understanding of microbial responses to antimicrobial agents. Time-lapse microscopy offers the advantage of visualizing antimicrobial effects on microbial cultures over time, capturing changes in cell morphology and growth behavior, and providing insights into the mode of action of antimicrobial agents. Although these cutting-edge methods are currently less commonly used for large-scale antimicrobial screening, they have the potential to significantly contribute to enhancing the understanding of antimicrobial compounds and furthering the fight against antimicrobial-resistant pathogens.
Conclusions and outlook
Antimicrobial research is witnessing remarkable growth as scientists strive to discover novel and potent antimicrobial agents to combat infectious diseases. This surge in research is fueled by the urgent need to address antimicrobial resistance and its impact on global health. In this pursuit, continuous efforts are being made to develop updated and efficient methods that can effectively identify and evaluate these potential antimicrobial agents. However, no single method can be universally deemed the best, as factors such as equipment availability, sample volume, and the nature of the agents being tested influence the choice of approach. Researchers are increasingly favoring less laborious, automated, faster, and user-friendly methods to streamline the antimicrobial screening process. While advanced technologies, such as high-throughput sequencing and advanced microscopy, hold promise in analyzing the mechanisms of action and detailed properties of antimicrobial agents, their widespread application and efficacy in antimicrobial assays are yet to be fully realized.
Efforts to combine multiple methods or complement existing ones are being explored to enhance the accuracy and reliability of antimicrobial screening. Ongoing improvements and updates to both manual and automated systems are crucial to ensure consistent and robust results. By tailoring method selection to specific needs and optimizing existing techniques accordingly, researchers can achieve a suitable equilibrium between simplicity and sensitivity in the assay methods. Ultimately, the adoption and refinement of these antimicrobial testing methods hold great promise in the discovery and development of effective antimicrobial agents. This progress contributes to a better management of infectious diseases and strengthens the fight against the growing threat of antimicrobial resistance, thus safeguarding public health worldwide.
Funding
This study received no fund from any organization.
Conflicts of interest
The author has no conflicts of interest to declare.
Ethical statement
The author declares that the research for the manuscript submitted does not involve human or animal research subjects.
Author contributions
TJH planned and wrote the manuscript.
Acknowledgment
TJH is grateful to his family members for invaluable supports while preparing this manuscript.
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