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I am working on optimizing growth rates for a culture, and one thing I'm trying to monitor is nutrient consumption rates to pinpoint any bottlenecks, i.e. the culture goes through all the calcium in a few hours and then stops growing.
Short of manually taking samples every few minutes and running individual tests for wide arrays of nutrients (minerals, amino acids, nitrogen compounds, etc.), is there a standard way of monitoring nutrient levels automatically in real-time? This is a personal research project, so unfortunately I don't have access to industrial bioreactors and the like.
Enzyme-based sensors for taking real-time measurements of growth substrates do exist. In brief: an enzyme degrades the substrate and thus changes the local redox state which is detected electronically (example glucose sensor). There's no guarantee that the substrates that you want to measure can be measured using such sensors. Also I think those sensors are fairly expensive and are tailored to specific conditions (ie it's not as simple as dipping the sensor into your culture medium). Practically speaking I think systems using these sensors are limited to monitoring just a small number of substrates.
LCMS or GCMS is the standard method for measuring concentrations of substrates. LC or GC (without the MS) may be possible, but probably not since there are are many components to culture media and so lots of overlapping peaks. Using this method is labor intensive (lots of samples), likely to be low resolution (temporally), and the equipment is pretty pricey, but if you have access to an LCMS and are willing to put the time in to become an LCMS expert then this is a good option.
Culture based methods using minimal media supplemented with substrates is an option, these experiments may achieve the same thing as real-time monitoring of substrate concentrations. If you have access to a plate reader this will be so much easier. The speed at which different substrates are consumed, and the order in which they are consumed can be inferred by measuring the growth rate of the bacteria and the final population size/dry weight of the culture.
I mention this culture-based method because, although it's not exactly what your question was about, it's much cheaper than the other options and much more established (and in my opinion more fun: you get a great feel for how the bacteria grow and what media they like, and the graphs look great).
Stream and River Monitoring
Without water, no life could exist, and many essential and nonessential human activities wouldn’t be possible without the use of healthy watersheds. These same activities can impact watersheds, in ways both large and small. Watersheds often span political and cultural boundaries while neighbors separated by city, state or national borders may not live under the same legal and cultural guidelines as one another, both could be citizens of the same watershed. By this measure, ensuring the health of a watershed — or the lakes, streams and rivers within — is as much a responsibility to your fellow human as it is to your local, state or federal regulating agency. For this same reason, water quality regulations are increasingly focused on the watershed level rather than established by political boundaries.
Streams and rivers offer an above ground glimpse at the health and hydrology of a watershed, and function as a vital resource for human activity, as well as habitat for a host of non-human animals and plants. In the U.S. alone, there are over 3.5 million miles of streams and rivers flowing through many different landscapes. Despite this widespread reliance on waterways, the U.S. Environmental Protection Agency has found that over half of streams and rivers in the U.S. are in poor biological condition. If a stream or river may be impacted by your project, it is vital to establish a proper monitoring system to ensure that the waterway’s hydrology and water quality are affected as little as possible, and so that any impact can be mitigated if it is detected.
Adenosine triphosphate (ATP) is the key energy source for all living organisms, essential to fundamental processes in all cells from metabolism to DNA replication and protein synthesis . In humans, abnormal cellular ATP levels and power consumption (ATP consumption rate), as can be determined by measuring and modeling ATP, are related to many diseases, such as cancer, aging, obesity, diabetes, neuronal disorders, viral infections, and immune dysfunctions [1,2,3,4,5,6]. In bacteria, ATP dynamics are directly related to bacterial metabolic activity, physiology, and behaviors under varying conditions and stresses [7,8,9]. For example, low ATP levels contribute to bacterial resistance/persistence in response to antibiotic treatments [9,10,11,12]. Despite ATP’s importance, our current understanding of ATP dynamics and homeostasis in cells has been limited by the lack of readily available and easy-to-use continuous ATP biosensors as well as by the shortage of accurate dynamic models to determine ATP fluxes.
The quantitative and continuous measurement of cellular ATP has proven challenging. Conventional methods, such as luciferase assays, require efficient lysis of cells and thus preclude real-time and continuous intracellular ATP measurements . To this end, several genetically encoded ATP biosensors have been developed, such as the fluorescence resonance energy transfer FRET-based ATeam biosensor , the bioluminescence resonance energy transfer BRET-based BTeam biosensor , and the new ATeam3.10 biosensor . These ratio-metric biosensors measure ATP, irrespective of their expression levels in the cell, and function well in slow-growing mammalian cell lines. To monitor cellular ATP in fast-growing bacteria, Yaginuma et al. developed a QUEEN ATP sensor but wider applications of this sensor in bacteria have not been reported, possibly due to its relatively dim signal and sensitivity to temperature . Furthermore, these biosensors require expensive fluorescence microscopes and time-consuming procedures for sample preparation and image analysis. These limitations make it challenging to continuously monitor intracellular ATP, e.g., in synthetic biological applications in the body that require fast, cheap, and continuous sensing of ATP in living microbial or other cells. Monitoring such ATP dynamics can predict nutrients, cellular stresses, disease states, or efficacy of drug treatments [2,3,4,5,6,7, 9, 11] and might be used to modulate or actuate therapeutic molecular release in response to cellular energetics.
Given that protein synthesis is the major energy-consuming process in the cell, ribosome synthesis must be tightly controlled by ATP/GTP availability in order to maintain ATP homeostasis [17,18,19,20]. The activity of a ribosomal RNA (rRNA) promoter, rrnB P1, has been shown to depend on cellular ATP level in E. coli [17, 18]. Upon binding, an RNA polymerase holoenzyme (RNAP) and the rrnB P1 promoter form a very short-lived open complex this unstable open complex requires an unusually high concentration of ATP (Kd in the mM range) to initiate the transcription of rRNA [17, 18, 21, 22]. The sensitivity of the rrnB P1 promoter to ATP is attributed to its specific features, including non-consensus -35 hexamers, non-optimal spacing between -35 and -10 hexamers, and a GC- rich discriminator [23, 24]. The requirement of high ATP concentration for transcription initiation is the rate-limiting step and allows for the regulation of rRNA production by changing ATP levels as long as they are not saturating [23, 24]. Therefore, the activity of the rrnB P1 promoter was proposed as a sensitive ATP indicator in E. coli [10, 17, 18]. However, systematic and quantitative analyses of rrnB P1-based ATP reporters that enable dynamic energy and power consumption measurements have thus far been missing. The combinatorial use of such ATP reporter and dynamic models could enable efficient determination of cellular energetics across various growth phases.
In this work, we designed and screened a series of synthetic ATP reporters in E. coli. The ATP reporters were made by fusing the ATP-sensing rrnB P1 promoter with the gene of a fast-folding GFP (GFP-mut2) that folds within minutes . An SsrA protease degradation tag [26, 27] fused to the C-terminus of the GFP also enabled its rapid degradation. Thus, the GFP produced from the rrnB P1 promoter in response to cellular ATP enabled relatively fast tracking of ATP in E. coli. We tested the performance of the reporter in minimal and rich media. Even though the activity of the rrnB P1 promoter is also affected by high levels of guanosine tetraphosphate (ppGpp) under starvation conditions [18, 20], we found that our ATP reporter can faithfully track cellular ATP levels under different experimental conditions regardless of potential ppGpp presence. After verifying the performance of the rrnB P1-based ATP reporter in various media and E. coli strains, we utilized it to study how bacterial ATP dynamics change in response to varying nutrients, including glucose and phosphate. To demonstrate the accuracy of the ATP reporter in power consumption measurements during bacterial growth, we developed a kinetic model and an electrical circuit model for bacterial growth in minimal medium. Our ATP reporter measurements and model enable us to quantitatively estimate intracellular ATP power consumption (ATP flux) in living cells, which is hard to estimate by luciferase-based or other ATP sensors. We show that our work can help quantify striking changes in ATP dynamics and power consumption across bacterial growth phases.
Nitrogen and Water
Nutrients, such as nitrogen and phosphorus, are essential for plant and animal growth and nourishment, but the overabundance of certain nutrients in water can cause several adverse health and ecological effects.
Nitrogen and Water
Sugar Creek, Indiana, is a creek running through fertilized farmland.
Nutrients, such as nitrogen and phosphorus, are essential for plant and animal growth and nourishment, but the overabundance of certain nutrients in water can cause a number of adverse health and ecological effects. Nitrogen, in the forms of nitrate, nitrite, or ammonium, is a nutrient needed for plant growth. About 78% of the air that we breathe is composed of nitrogen gas, and in some areas of the United States, particularly the northeast, certain forms of nitrogen are commonly deposited in acid rain.
Of course, nitrogen is used in agriculture to grow crops, and on many farms the landscape has been greatly modified to maximize farming output. Fields have been leveled and modified to efficiently drain off excess water that may fall as precipitation or from irrigation practices.
This image shows Sugar Creek in Indiana, as it has been extensively modified for human use. As commonly found in small agricultural streams, Sugar Creek has been straightened, deepened, and had tile drains installed to favor rapid removal of water from agricultural lands. If excess nitrogen is found in the crop fields, the drainage water can introduce it into streams like these, which will drain into other larger rivers and might end up in the Gulf of Mexico, where excess nitrogen can lead to hypoxic conditions (lack of oxygen).
Sources of nitrogen
Fertilizers and other chemicals are applied to crop fields worldwide. Due to runoff, excess chemicals can find their way into water bodies and harm water quality.
Credit: Pixabay - Creative Commons
Although nitrogen is abundant naturally in the environment, it is also introduced through sewage and fertilizers. Chemical fertilizers or animal manure is commonly applied to crops to add nutrients. It may be difficult or expensive to retain on site all nitrogen brought on to farms for feed or fertilizer and generated by animal manure. Unless specialized structures have been built on the farms, heavy rains can generate runoff containing these materials into nearby streams and lakes. Wastewater-treatment facilities that do not specifically remove nitrogen can also lead to excess levels of nitrogen in surface or groundwater.
Nitrate can get into water directly as the result of runoff of fertilizers containing nitrate. Some nitrate enters water from the atmosphere, which carries nitrogen-containing compounds derived from automobiles and other sources. More than 3 million tons of nitrogen are deposited in the United States each year from the atmosphere, derived either naturally from chemical reactions or from the combustion of fossil fuels, such as coal and gasoline. Nitrate can also be formed in water bodies through the oxidation of other forms of nitrogen, including nitrite, ammonia, and organic nitrogen compounds such as amino acids. Ammonia and organic nitrogen can enter water through sewage effluent and runoff from land where manure has been applied or stored.
Sources of nitrogen to the Gulf of Mexico
Identifying nutrient sources is a complicated task because, at more than 1.2 million square miles, the Mississippi River Basin is the fourth largest basin in the world. It covers close to 40 percent of the lower 48 States. There are 31 States that drain, via the Mississippi River Basin, into the Gulf of Mexico, and nutrient sources are found throughout the basin.
Fertilizers used on crops, air pollution, and manure are some of the major sources of nitrogen transported from the Mississippi River Basin to the Gulf of Mexico.
Problems with excess levels of nitrogen in the environment
Excess nitrogen can harm water bodies
Excess nitrogen can cause overstimulation of growth of aquatic plants and algae. Excessive growth of these organisms, in turn, can clog water intakes, use up dissolved oxygen as they decompose, and block light to deeper waters. Lake and reservoir eutrophication can occur, which produces unsightly scums of algae on the water surface, can occasionally result in fish kills, and can even "kill" a lake by depriving it of oxygen. The respiration efficiency of fish and aquatic invertebrates can occur, leading to a decrease in animal and plant diversity, and affects our use of the water for fishing, swimming, and boating.
Excess nitrogen in water can harm people
An algae bloom on Lake Le-Auqa-Na, Illinois.
Too much nitrogen, as nitrate, in drinking water can be harmful to young infants or young livestock. Excessive nitrate can result in restriction of oxygen transport in the bloodstream. Infants under the age of 4 months lack the enzyme necessary to correct this condition ("blue baby syndrome").
Variation of nitrate across the United States
The concentration of nitrate (a form of nitrogen) of water bodies vary widely across the United States. Natural and human processes determine concentration of nitrate in water. The National Atmospheric Deposition Program has developed maps showing nitrate patterns, such as the one below showing the spatial pattern of nitrate at selected sampling sites for 2002. You should be aware that this contour map was developed using the nitrate measurements at the specific sampling locations thus, the contours and isolines were created using interpolation between data points. You should not necessarily use the map to document the nitrate of a water body at a particular map location, but rather, use the map as a general indicator of nitrate throughout the country.
Source: National Atmospheric Deposition Program (NRSP-3)/National Trends Network. (2004). NADP Program Office, Illinois State Water Survey, 2204 Griffith Dr., Champaign, IL 61820.
Risks of nitrate contamination in shallow groundwater
Much of the Nation uses groundwater at its main source of water for many needs, from drinking water and other home uses to irrigation to public uses, such as supplying water to parks. Of course, geology and the factors that affect the availability of groundwater vary greatly geographically, but many places, such as southern Georgia, have aquifers that can supply a lot of freshwater very near the land surface. Since nitrogen contamination is more of a problem in shallow aquifers, it is worthwhile to be aware of what aquifers in the United States would be more at risk for nitrogen contamination.
Developed for a USGS study, the map below shows those areas with the highest risk for contamination of shallow groundwater by nitrate. Generally, aquifer vulnerability is represented by soil-drainage characteristics—the ease with which water and chemicals can seep to groundwater—and the extent to which woodlands are interspersed with crop land. Use of the risk map to identify and prioritize contamination at a more detailed level than presented here is not advised because local variations in land use, irrigation practices, aquifer type, and rainfall can result in nitrate concentrations that do not conform to risk patterns shown at the national scale.
This chapter is an introductory overview of the most commonly used assay methods to estimate the number of viable cells in multi-well plates. This chapter describes assays where data are recorded using a plate-reader it does not cover assay methods designed for flow cytometry or high content imaging. The assay methods covered include the use of different classes of colorimetric tetrazolium reagents, resazurin reduction and protease substrates generating a fluorescent signal, the luminogenic ATP assay, and a novel real-time assay to monitor live cells for days in culture. The assays described are based on measurement of a marker activity associated with viable cell number. These assays are used for measuring the results of cell proliferation, testing for cytotoxic effects of compounds, and for multiplexing as an internal control to determine viable cell number during other cell-based assays.
Cell-based assays are often used for screening collections of compounds to determine if the test molecules have effects on cell proliferation or show direct cytotoxic effects that eventually lead to cell death. Cell-based assays also are widely used for measuring receptor binding and a variety of signal transduction events that may involve the expression of genetic reporters, trafficking of cellular components, or monitoring organelle function. Regardless of the type of cell-based assay being used, it is important to know how many viable cells are remaining at the end of the experiment. There are a variety of assay methods that can be used to estimate the number of viable eukaryotic cells. This chapter will provide an overview of some of the major methods used in multi-well formats where data are recorded using a plate reader. The methods described include: tetrazolium reduction, resazurin reduction, protease markers, and ATP detection. Methods for flow cytometry and high content imaging may be covered in different chapters in the future.
The tetrazolium reduction, resazurin reduction, and protease activity assays measure some aspect of general metabolism or an enzymatic activity as a marker of viable cells. All of these assays require incubation of a reagent with a population of viable cells to convert a substrate to a colored or fluorescent product that can be detected with a plate reader. Under most standard culture conditions, incubation of the substrate with viable cells will result in generating a signal that is proportional to the number of viable cells present. When cells die, they rapidly lose the ability to convert the substrate to product. That difference provides the basis for many of the commonly used cell viability assays. The ATP assay is somewhat different in that the addition of assay reagent immediately ruptures the cells, thus there is no incubation period of reagent with a viable cell population.
Tetrazolium Reduction Assays
A variety of tetrazolium compounds have been used to detect viable cells. The most commonly used compounds include: MTT, MTS, XTT, and WST-1. These compounds fall into two basic categories: 1) MTT which is positively charged and readily penetrates viable eukaryotic cells and 2) those such as MTS, XTT, and WST-1 which are negatively charged and do not readily penetrate cells. The latter class (MTS, XTT, WST-1) are typically used with an intermediate electron acceptor that can transfer electrons from the cytoplasm or plasma membrane to facilitate the reduction of the tetrazolium into the colored formazan product.
MTT Tetrazolium Assay Concept
The MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) tetrazolium reduction assay was the first homogeneous cell viability assay developed for a 96-well format that was suitable for high throughput screening (HTS) (1). The MTT tetrazolium assay technology has been widely adopted and remains popular in academic labs as evidenced by thousands of published articles. The MTT substrate is prepared in a physiologically balanced solution, added to cells in culture, usually at a final concentration of 0.2 - 0.5mg/ml, and incubated for 1 to 4 hours. The quantity of formazan (presumably directly proportional to the number of viable cells) is measured by recording changes in absorbance at 570 nm using a plate reading spectrophotometer. A reference wavelength of 630 nm is sometimes used, but not necessary for most assay conditions.
Viable cells with active metabolism convert MTT into a purple colored formazan product with an absorbance maximum near 570 nm (Figure 1). When cells die, they lose the ability to convert MTT into formazan, thus color formation serves as a useful and convenient marker of only the viable cells. The exact cellular mechanism of MTT reduction into formazan is not well understood, but likely involves reaction with NADH or similar reducing molecules that transfer electrons to MTT (2). Speculation in the early literature involving specific mitochondrial enzymes has led to the assumption mentioned in numerous publications that MTT is measuring mitochondrial activity (3, 4).
Structures of MTT and colored formazan product.
The formazan product of the MTT tetrazolium accumulates as an insoluble precipitate inside cells as well as being deposited near the cell surface and in the culture medium. The formazan must be solubilized prior to recording absorbance readings. A variety of methods have been used to solubilize the formazan product, stabilize the color, avoid evaporation, and reduce interference by phenol red and other culture medium components (5-7). Various solubilization methods include using: acidified isopropanol, DMSO, dimethylformamide, SDS, and combinations of detergent and organic solvent (1, 5-7). Acidification of the solubilizing solution has the benefit of changing the color of phenol red to yellow color that may have less interference with absorbance readings. The pH of the solubilization solution can be adjusted to provide maximum absorbance if sensitivity is an issue (8) however, other assay technologies offer much greater sensitivity than MTT.
The amount of signal generated is dependent on several parameters including: the concentration of MTT, the length of the incubation period, the number of viable cells and their metabolic activity. All of these parameters should be considered when optimizing the assay conditions to generate a sufficient amount of product that can be detected above background.
The conversion of MTT to formazan by cells in culture is time dependent (Figure 2).
Direct correlation of formazan absorbance with B9 hybridoma cell number and time-dependent increase in absorbance. Note: there is little absorbance change between 2 and 4 hours. Adapted from CellTiter 96 ® Non-Radioactive Cell Proliferation Assay (more. )
Longer incubation time will result in accumulation of color and increased sensitivity up to a point however, the incubation time is limited because of the cytotoxic nature of the detection reagents which utilize energy (reducing equivalents such as NADH) from the cell to generate a signal. For cell populations in log phase growth, the amount of formazan product is generally proportional to the number of metabolically active viable cells as demonstrated by the linearity of response in Figure 2. Culture conditions that alter the metabolism of the cells will likely affect the rate of MTT reduction into formazan. For example, when adherent cells in culture approach confluence and growth becomes contact inhibited, metabolism may slow down and the amount MTT reduction per cell will be lower. That situation will lead to a loss of linearity between absorbance and cell number. Other adverse culture conditions such as altered pH or depletion of essential nutrients such as glucose may lead to a change in the ability of cells to reduce MTT.
The MTT assay was developed as a non-radioactive alternative to tritiated thymidine incorporation into DNA for measuring cell proliferation (1). In many experimental situations, the MTT assay can directly substitute for the tritiated thymidine incorporation assay (Figure 3).
A comparison of using the MTT and 3 [H]thymidine incorporation assays of hGM-CSF-treated TF-1 cells. A blank absorbance value of 0.065 (from wells without cells but treated with MTT) was subtracted from all absorbance values. Adapted from CellTiter 96 (more. )
However, it is worth noting that MTT reduction is a marker reflecting viable cell metabolism and not specifically cell proliferation. Tetrazolium reduction assays are often erroneously described as measuring cell proliferation without the use of proper controls to confirm effects on metabolism (10).
Shortly after addition of MTT, the morphology of some cell types can be observed to change dramatically suggesting altered physiology (11 and Figure 4).
Change in NIH3T3 cell morphology after exposure to MTT (0.5 mg/ml). Panel A shows a field of cells photographed immediately after addition of the MTT solution. Panel B shows the same field of cells photographed after 4 hours of exposure to MTT. Panel (more. )
Toxicity of the MTT compound is likely related to the concentration added to cells. Optimizing the concentration may result in lower toxicity. Given the cytotoxic nature of MTT, the assay method must be considered as an endpoint assay. A recent report speculated that formazan crystals contribute to harming cells by puncturing membranes during exocytosis (12). The observation of extracellular formazan crystals many times the diameter of cells that grow longer over time make it seem unlikely that exocytosis of those large structures was involved (Figure 4 and 5).
U937 cells incubated with MMT tetrazolium for 3 hours showing formazan crystals larger than the cells. Image was captured using an Olympus FV500 confocal microscope. Scale bar = 20 µm
Growing crystals may suggest that marginally soluble formazan accumulates where seed crystals have begun to deposit.
Reducing compounds are known to interfere with tetrazolium reduction assays. Chemicals such as ascorbic acid, or sulfhydryl-containing compounds including reduced glutathione, coenzyme A, and dithiothreitol, can reduce tetrazolium salts non-enzymatically and lead to increased absorbance values in assay wells (13-17). Culture medium at elevated pH or extended exposure of reagents to direct light also may cause an accelerated spontaneous reduction of tetrazolium salts and result in increased background absorbance values. Suspected chemical interference of test compounds can be confirmed by measuring absorbance values from control wells without cells incubated with culture medium containing MTT and various concentrations of the test compound.
Commercial kits containing solutions of MTT and a solubilization reagent as well as MTT reagent powder are available from several vendors. For example:
The concentration of the MTT solution and the nature of the solubilization reagent differ among various vendors. The amount of formazan signal generated will depend on variety of parameters including the cell type, number of cells per well, culture medium, etc. Although the commercially available kits are broadly applicable to a large number of cell types and assay conditions, the concentration of the MTT and the type of solubilization solution may need to be adjusted for optimal performance.
Dissolve MTT in Dulbecco’s Phosphate Buffered Saline, pH=7.4 (DPBS) to 5 mg/ml.
Filter-sterilize the MTT solution through a 0.2 µM filter into a sterile, light protected container.
Store the MTT solution, protected from light, at 4ଌ for frequent use or at -20ଌ for long term storage.
Choose appropriate solvent resistant container and work in a ventilated fume hood.
Prepare 40% (vol/vol) dimethylformamide (DMF) in 2% (vol/vol) glacial acetic acid.
Add 16% (wt/vol) sodium dodecyl sulfate (SDS) and dissolve.
Store at room temperature to avoid precipitation of SDS. If a precipitate forms, warm to 37ଌ and mix to solubilize SDS.
MTT Assay Protocol
Prepare cells and test compounds in 96-well plates containing a final volume of 100 µl/well.
Incubate for desired period of exposure.
Add 10 µl MTT Solution per well to achieve a final concentration of 0.45 mg/ml.
Incubate 1 to 4 hours at 37ଌ.
Add 100 µl Solubilization solution to each well to dissolve formazan crystals.
Mix to ensure complete solubilization.
Record absorbance at 570 nm.
MTS Tetrazolium Assay Concept
More recently developed tetrazolium reagents can be reduced by viable cells to generate formazan products that are directly soluble in cell culture medium. Tetrazolium compounds fitting this category include MTS, XTT, and the WST series (18-23). These improved tetrazolium reagents eliminate a liquid handling step during the assay procedure because a second addition of reagent to the assay plate is not needed to solubilize formazan precipitates, thus making the protocols more convenient. The negative charge of the formazan products that contribute to solubility in cell culture medium are thought to limit cell permeability of the tetrazolium (24). This set of tetrazolium reagents is used in combination with intermediate electron acceptor reagents such as phenazine methyl sulfate (PMS) or phenazine ethyl sulfate (PES) which can penetrate viable cells, become reduced in the cytoplasm or at the cell surface and exit the cells where they can convert the tetrazolium to the soluble formazan product (25). The general reaction scheme for this class of tetrazolium reagents is shown in Figure 6.
Intermediate electron acceptor pheazine ethyl sylfate (PES) transfers electron from NADH in the cytoplasm to reduce MTS in the culture medium into an aqueous soluble formazan.
In general, this class of tetrazolium compounds is prepared at 1 to 2mg/ml concentration because they are not as soluble as MTT. The type and concentration of the intermediate electron acceptor used varies among commercially available reagents and in many products the identity of the intermediate electron acceptor is not disclosed. Because of the potential toxic nature of the intermediate electron acceptors, optimization may be advisable for different cell types and individual assay conditions. There may be a narrow range of concentrations of intermediate electron acceptor that result in optimal performance.
Commercial kits containing solutions of MTS, XTT, and WST-1 and an intermediate electron acceptor reagent are available from several vendors. For example:
Nutrient pollution is one of America's most widespread, costly and challenging environmental problems, and is caused by excess nitrogen and phosphorus in the air and water.
Nitrogen and phosphorus are nutrients that are natural parts of aquatic ecosystems. Nitrogen is also the most abundant element in the air we breathe. Nitrogen and phosphorus support the growth of algae and aquatic plants, which provide food and habitat for fish, shellfish and smaller organisms that live in water.
But when too much nitrogen and phosphorus enter the environment - usually from a wide range of human activities - the air and water can become polluted. Nutrient pollution has impacted many streams, rivers, lakes, bays and coastal waters for the past several decades, resulting in serious environmental and human health issues, and impacting the economy.
Too much nitrogen and phosphorus in the water causes algae to grow faster than ecosystems can handle. Significant increases in algae harm water quality, food resources and habitats, and decrease the oxygen that fish and other aquatic life need to survive. Large growths of algae are called algal blooms and they can severely reduce or eliminate oxygen in the water, leading to illnesses in fish and the death of large numbers of fish. Some algal blooms are harmful to humans because they produce elevated toxins and bacterial growth that can make people sick if they come into contact with polluted water, consume tainted fish or shellfish, or drink contaminated water.
Nutrient pollution in ground water - which millions of people in the United States use as their drinking water source - can be harmful, even at low levels. Infants are vulnerable to a nitrogen-based compound called nitrates in drinking water. Excess nitrogen in the atmosphere can produce pollutants such as ammonia and ozone, which can impair our ability to breathe, limit visibility and alter plant growth. When excess nitrogen comes back to earth from the atmosphere, it can harm the health of forests, soils and waterways.
Preserving Bacterial Cultures
Bacteria can be stored for months or years if they are stored at -80C and in a high percentage of glycerol.
Describe how bacterial cultures can be stored for a long time at -80C in glycerol
- Preserve your selected bacteria so you always have something to go back to if something goes wrong.
- While it is possible to make a long term stock from cells in stationary phase, ideally your culture should be in logarithmic growth phase.
- Ensure a pure culture is being preserved by picking a single colony of the bacteria off a plate for cryopreservation.
- cryogenic: of, relating to, or performed at low temperatures
- logarithmic growth phase: exponential phase (sometimes called the log phase or the logarithmic phase) is a period characterized by cell doubling.
Bacteria in liquid media: An erlenmeyer containing a bacterial culture. Bacteria that have been preserved in glycerol stocks can be grown overnight in liquid media to promote propagation.
Three species of bacteria, Carnobacterium pleistocenium, Chryseobacterium greenlandensis, and Herminiimonas glaciei, have reportedly been revived after surviving for thousands of years frozen in ice. As a practical matter, as a researcher, you will want to preserve your selected bacteria so you can go back to it if something goes wrong.
Whenever you successfully transform a bacterial culture with a plasmid or whenever you obtain a new bacterial strain, you will want to make a long term stock of that bacteria. Bacteria can be stored for months and years if they are stored at -80C and in a high percentage of glycerol.
In order to ensure a pure culture is being preserved, pick a single colony of the bacteria off a plate, grow it overnight in the appropriate liquid media, and with shaking. Take the overnight culture and and mix an aliquot with 40% glycerol in sterile water and place in a cryogenic vial. It is important to label the vial with all the relevant information (e.g. strain, vector, date, researcher, etc.). Freeze the glycerol stock and store at -80C. At this point you should also record the strain information and record the location.
While it is possible to make a long term stock from cells in the stationary phase, ideally your culture should be in logarithmic growth phase. Certain antibiotics in the medium should be removed first as they are supposedly toxic over time, e.g. Tetracycline. To do this, spin the culture down and resuspend it in the same volume of straight LB medium.
Food Calorimetry: How to Measure Calories in Food
People who check nutrition labels to make informed decisions about which foods to eat and which to avoid often base those decisions solely on the number of calories per serving. A calorie, like a joule, is a unit of energy. The International System of Units (SI) unit for energy is the joule however, the calorie is commonly used for a unit of food energy. A calorie is equal to the amount of energy per unit mass required to raise the temperature of 1 g of water by 1° C. One calorie is the equivalent of 4.18 joules. Food calories, as read off a nutrition label, are actually kilocalories (often denoted as lories” with a capital C). There are 1,000 calories in a kilocalorie, or food Calorie.
A calorimeter is a piece of equipment designed to measure the energy released or absorbed during a chemical reaction or phase change. Food calorimetry allows us to determine the number of calories per gram of food. In this activity, a piece of food is burned and the released energy is used to heat a known quantity of water. The temperature change (∆T) of the water is then used to determine the amount of energy in the food.
Use safety glasses or goggles and be cautious with the matches and burning food samples. Check for food allergies before using food samples. Sensitive individuals should not participate in any activities that may result in exposure. Never eat or drink in lab.
- Soda Can (empty)
- Stirring Rod
- Ring Stand and Ring
- Graduated Cylinder, 100 mL
- Large Paper Clip
- 2 Twist Ties
- 3 Food Samples with Nutrition Labels (2 to 3 g each of samples such as nuts marshmallows or soft chips, e.g., cheese puffs)
- Aluminum Foil (small piece)
At least 1 of the following to be shared
- Using the graduated cylinder, obtain 50 mL of water and carefully pour it into the soda can.
- Determine the mass of water and record your finding in the data table (hint: density of water = 1 g/mL).
- Hold the paper clip horizontally and bend the outer end upwards until it is at a 90° angle to the rest of the paper clip.
- Obtain a 2- to 3-g food sample.
- Place the food sample on the paper clip&aposs upward-extending end. The sample should be freestanding, supported by the bottom of the paper clip (see Fig. 1). Determine the initial mass of the food sample and paper clip, and record your findings in the data table.
- Place a small piece of aluminum foil underneath the paper clip in a space that has been cleared of all flammables.
- Insert the stirring rod through the soda can tab and position the can in the ring stand so the stirring rod supports it (see Fig. 2).
- Adjust the ring stand until the can is approximately 4 cm above the food sample.
- Suspend the thermometer inside the can a few centimeters above the can&aposs bottom. Secure with 2 twist ties.
- Determine the initial temperature of the water in the can and record this value in the data table.
- Carefully light a match and use it to light the food sample.
- Allow the lit sample to heat the water in the can. Gently stir the water periodically with the thermometer (see Fig. 3).
- Monitor the temperature change of the water and record the highest observed temperature in the data table.
Eukaryotes have two major protein degradation systems within cells. One is the ubiquitin-proteasome system, which accounts for the selective degradation of most short-lived proteins (Hochstrasser, 1996 Hershko and Ciechanover, 1998). The other is the lysosomal system. Proteins from both inside and outside of the cell are delivered to the lytic compartment. Degradation of exogenous materials and plasma membrane proteins is mediated by the process of endocytosis/phagocytosis, whereas degradation of cytoplasmic components is achieved by autophagy (also known as autophagocytosis). Three types of autophagy have been proposed: macroautophagy, microautophagy, and chaperon-mediated autophagy (Seglen and Bohley, 1992 Dunn, 1994 Blommaart et al., 1997). Macroautophagy is thought to be responsible for the majority of the intracellular protein degradation in mammalian cells, particularly during starvation-induced proteolysis (Mortimore and Pösö, 1987).
Macroautophagy (simply referred to as autophagy hereafter) is mediated by a unique organelle termed the autophagosome. A membrane cisterna called the isolation membrane (also known as phagophore) encloses a portion of cytoplasm, resulting in the formation of the autophagosome. The autophagosome is a double-membrane structure containing undigested cytoplasmic materials including organelles. The sequestration step is generally thought to be nonselective. Next, the outer membrane of the autophagosome fuses with the lysosome membrane. Various hydrolytic enzymes are supplied to the autophagosome and the cytoplasm-derived contents are degraded together with the inner membrane of the autophagosome. This degrading structure is termed the autolysosome/autophagolysosome.
Autophagy is thought to be required for normal turnover of cellular components particularly in response to starvation (Mortimore and Pösö, 1987). Autophagy-defective yeast cells die quickly during starvation (Tsukada and Ohsumi, 1993). Autophagy also plays an important role in some types of differentiation/development: ATG genes (described below) are essential for spore formation in yeast (Tsukada and Ohsumi, 1993), and the development of Drosophila melanogaster (Juhasz et al., 2003), Dictyostelium discoideum (Otto et al., 2003), and Caenorhabditis elegans (Melendez et al., 2003). Plants deficient for autophagy genes show acceleration of senescence (Doelling et al., 2002 Hanaoka et al., 2002). In contrast, the precise roles of autophagy in mammals are not known, although a growing number of studies have suggested that autophagy might be important for cell death during embryogenesis (Clarke, 1990) and pathogenesis (Liang et al., 1999 Nishino et al., 2000). In addition, systematic analysis describing where and when autophagy occurs has not been performed. This is largely due to a lack of good diagnostic methods. To date, electron microscopy has been the only method to monitor autophagy. Unfortunately, this is a method requiring many skills and much time, and sometimes it is difficult to distinguish autophagic vacuoles from other structures just by morphology. Although an elegant transgenic mouse model was recently generated to monitor the ubiquitin/proteasome system (Lindsten et al., 2003), we have not had such in vivo assay systems for autophagy.
We have dissected the autophagic pathway at the molecular level using both yeast and mammalian cells. In the yeast, Saccharomyces cerevisiae, at least 16 of APG and AUT genes have been identified to be required for autophagosome formation (Klionsky and Ohsumi, 1999 Ohsumi, 2001 Mizushima et al., 2002a). The nomenclature of these autophagy-related genes were recently unified as ATG (Klionsky et al., 2003). We have found two novel ubiquitylation-like conjugation systems: one mediates conjugation of Atg12 to Atg5 (Mizushima et al., 1998a) and the other mediates a covalent linkage between Atg8 (Aut7/Atg8) and phosphatidylethanolamine (PE Ichimura et al., 2000). The resulting conjugates, Atg12-Atg5 and Atg8-PE, function in autophagosome formation (Suzuki et al., 2001 Noda et al., 2002). These two conjugation systems are highly conserved in mammals (Mizushima et al.,1998b, 2002b Kabeya et al., 2000 Tanida et al.,2001, 2002). Most Atg12-Atg5 conjugate exists in the cytosol as a complex with Atg16L (Mizushima et al.,1999, 2003 Kuma et al., 2002). Only a small fraction of the Atg12-Atg5·Atg16L complex localizes to the autophagic isolation membranes throughout its elongation process, and the complex dissociates from the membrane when autophagosome formation is completed (Mizushima et al., 2001). We have demonstrated that mouse Atg12-Atg5 conjugate is indeed essential for the membrane elongation process by generating ATG5 -/- embryonic stem (ES) cells (Mizushima et al., 2001). LC3, one of the mammalian homologues of Atg8, also targets the isolation membrane in an Atg5-dependent manner. However, LC3 remains on the membrane even after spherical autophagosomes are completely formed (Kabeya et al., 2000 Mizushima et al., 2001). When we label autophagosomes with green fluorescent protein (GFP)-LC3, we are able to see them by fluorescence microscopy as ring-shaped structures (Mizushima et al., 2003), or dots in the case of small autophagosomes (Kabeya et al., 2000). Because autolysosomes have less LC3 on the membrane than autophagosomes, the fluorescent signal of autolysosomes is weaker. On the basis of these observations, we propose that LC3 could serve as a good molecular marker for isolation membranes, autophagosomes, and some autolysosomes (Kabeya et al., 2000).
We have applied this method to in vivo studies to monitor autophagy simply and accurately. We generated GFP-LC3 transgenic mice and examined the occurrence of autophagy in various tissues. Using this novel assay system, we performed comprehensive and quantitative analysis on the autophagic response during starvation.
Freshwater and marine algae can balance nutrient demand and availability by regulating uptake, accumulation and exudation. To obtain insight into these processes under nitrogen (N) and phosphorus (P) limitation, we reanalyze published data from continuous cultures of the chlorophyte Selenastrum minutum. Based on mass budgets, we argue that much of the non‐limiting N and P had passed through the organisms and was present as dissolved organic phosphorus or nitrogen (DOP or DON). We construct a model that describes the production of biomass and dissolved organic matter (DOM) as a function of the growth rate. A fit of this model against the chemostat data suggests a high turnover of the non‐limiting N and P: at the highest growth rates, N and P atoms spent on average only about 3 h inside an organism, before they were exuded as DON and DOP, respectively. This DOM exudation can explain the observed trends in the algal stoichiometric ratios as a function of the dilution rate. We discuss independent evidence from isotope experiments for this apparently wasteful behavior and we suggest experiments to quantify and characterize DON and DOP exudation further.