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Why is absorbance at 280 nm for protein solution going up when I measure repeatedly?

Why is absorbance at 280 nm for protein solution going up when I measure repeatedly?


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I have been measuring my protein solutions' concentrations by diluting them in water 20 fold with a final volume of 100 uL and then measuring the absorbances of these solutions in 96 well plates with plate reader. I don't remember having any problem up until today.

I used 20 mM phosphate buffer instead of water to dilute them today and measured the absorbances at 280 nm repeatedly three times and the absorbance for one of the solutions went up from 0.043 to 0.068 (absorbance of 20 mM phosphate buffer is 0.030 at 280 nM with same volume); I stopped measuring after third one but it would probably go higher as I measured until I hit a plateau.

I measured absorbances of two proteins and only one of them went up that much, other one went up from 0.071 to 0.088; if this were to be concentration dependent I would expect the second solution go even higher but it didn't happen.

I know there may be differences in UV absorbances if protein is folded or unfolded; would it be that dramatic? What is the reason for that increase? I will be grateful for an explanation and a practical solution to the problem.

NOTE: I increased the total volume to 200 uL by simply adding 100 uL of 20 mM phosphate buffer into all wells and the signal increase slowed down a lot; there is still some increase though with 0.001-0.003 increments in each measurement.


It looks like your protein concentrations are right on the limit of detection of the spectrophotometer, and changing the diluent buffer changed their concentrations. The samples may not have been thoroughly mixed after dilution and before measurement, so the varying measurements may simply be the solution coming to equilibrium. Temperature can also affect absorbance, so you should verify that your samples have equilibrated before drawing any conclusions. If the absorbance of your phosphate buffer is 0.03, I'd try to keep the sample absorbances above 0.075 or higher to avoid getting too close to the limit of detection. Also, make sure your buffer isn't too old or contaminated with something which could be affecting its absorbance characteristics.

I would suggest taking one or two protein samples and doing a dilution series (1:1, 1:5, 1:10, 1:20) in a large-ish volume (say 400 ul each, if you can spare it), vortex briefly to mix well, then measure triplicates of each dilution on your reader, along with appropriate blanks (buffer only). You will see differences between each measurement, but it should be quite small, depending on the accuracy and precision of your instrument.

Measured values will not be exactly the same from measurement to measurement, and it would take a lot more than three repetitions to determine if there was an actual drift trend occurring. Measure your sample plate every 5 minutes for an hour and plot the values (don't just eyeball them) to see if the machine may need to be serviced.


Could be protein unfolding or changes in conformation. Absorbance at 280 nm is mainly due to the tryptophan residues, and can change substantially as these residues move from a more hydrophobic (buried inside the protein) to a more hydrophilic (exposed to the solvent) environment. Your protein may be reacting to the new buffer.


  • Path length l is usually in units of cm. (note: most spectrophotometers are designed to accept 1cm wide cuvettes)
  • Molar extinction coefficient&epsilon has units of M -1 cm -1 and is a constant of proportionality that relates the absorption of molar solutions
  • Mass extinction coefficient&epsilon 1% refers to the absorbance of a 1% by mass solution. Typically this refers to an aqueous solution that we can take to have a density of 1000g/L. A 1% by mass aqueous solution would therefore refer to the dissolution of 10g/L, or a 10mg/ml solution of the molecule of interest.
  • Since the absorbance of a molecule is a function of the wavelength (i.e. the absorption is not equal for every wavelength) the extinction coefficient must also reference a wavelength. This is typically done using a subscript:

&epsilon 1% 280nm = 14.5 g -1 L cm -1

· In this case a 10mg/ml solution of the molecule will have an absorbance reading of 14.5 (dimensionless units) at l = 280nm (the absorption at other wavelengths may not be known). The units of concentration are g/L, thus e will have dimensions of g -1 L cm -1 .

Why is it important to be able to quantitate protein concentration in a sample?

An important application of "Biotechnology" is the production of proteins as commercial products. Such products might have pharmaceutical applications (e.g. insulin, human growth hormone, tissue plasminogen activator, erythropoietin, blood clotting factor VIII.), industrial applications (e.g. subtilisin (an enzyme in detergents), 2,5-diketo-D-gluconate reductase (an enzyme in vitamin C production), as materials (e.g. silk protein in textiles, barnacle adhesion protein as a glue). In these cases, there are various aspects of successful production that require quantitation:

  • How much of the protein can be produced (i.e. what is the efficiency of production)?
  • How pure is the protein that is produced (industrial applications may require 90% pure, pharmaceutical applications may require 99.999% pure)

Such proteins may be isolated from natural sources (e.g. blood clotting factor VIII may be extracted from human blood), or they may be produced recombinantly (e.g. E. coli bacterial cells can be genetically engineered to produce human growth hormone). In both cases, it may be necessary to purify the protein using a series of fractionation steps. We will go into more detail about such fractionation steps in a later lecture, but the general idea is that a heterogeneous mixture of molecules can be fractionated based upon some physical property of the molecules. The following are properties that can be used to fractionate a heterogeneous mixture of biomolecules:

  • Molecular mass (i.e. "big" molecules can be separated from "small" molecules)
  • pKa (i.e. "acidic" molecules can be separated from "basic" molecules)
  • Hydrophobicity (i.e. non-polar molecules can be separated from polar molecules)

For such fractionation steps involving proteins, we need to keep track of how much of the contaminating proteins went into one fraction and how much of our desired protein went into the other fraction. Although the details are somewhat more complicated than this simple description, it is important to be able to quantitate protein concentration to be able to effectively purify a protein of interest.

Once a protein is pure, it may be of considerable economic interest to be able to quantify the yield (and, therefore, be able to determine how much it cost to produce a given mass of protein). For example, the only source for human growth hormone (to treat small stature) used to be to extract it from human pituitary glands harvested from the brains of cadavers. Suffice it to say, this made the protein extremely expensive. Furthermore, the isolation from human tissues meant that the sample could also be potentially contaminated with human pathogens (hepatitis, CJD, AIDS, etc.). With the advent of genetic engineering, the production of human growth hormone by bacterial cells (i.e. E. coli) meant that relative large quantities could be produced far cheaper (and with no threat of human pathogens).

Why not just weigh the protein?

  • Most samples are typically quantities of milligrams or even micrograms, not grams, and thus, it is difficult to transfer and measure such small amounts
  • Water is present in proteins, and it is extremely difficult to remove all the water (some water molecules hydrogen bond extremely tightly to proteins). Thus, the mass measurement would include some waters, and would increase the apparent mass of the protein

The Absorbance of a Solution

For each wavelength of light passing through the spectrometer, the intensity of the light passing through the reference cell is measured. This is usually referred to as (I_o) - that's (I) for Intensity.

Figure (PageIndex<1>): Light absorbed by sample in a cuvette

The intensity of the light passing through the sample cell is also measured for that wavelength - given the symbol, (I). If (I) is less than (I_o), then the sample has absorbed some of the light (neglecting reflection of light off the cuvette surface). A simple bit of math is then done in the computer to convert this into something called the absorbance of the sample - given the symbol, (A). The absorbance of a transition depends on two external assumptions.

  1. The absorbance is directly proportional to the concentration ((c)) of the solution of the sample used in the experiment.
  2. The absorbance is directly proportional to the length of the light path ((l)), which is equal to the width of the cuvette.

Assumption one relates the absorbance to concentration and can be expressed as

The absorbance ((A)) is defined via the incident intensity (I_o) and transmitted intensity (I) by

Assumption two can be expressed as

Combining Equations ( ef<1>) and ( ef<3>):

This proportionality can be converted into an equality by including a proportionality constant ((epsilon)).

This formula is the common form of the Beer-Lambert Law, although it can be also written in terms of intensities:

[ A=log_ <10>left( dfrac ight) = epsilon l c label <6>]

The constant (epsilon) is called molar absorptivity or molar extinction coefficient and is a measure of the probability of the electronic transition. On most of the diagrams you will come across, the absorbance ranges from 0 to 1, but it can go higher than that. An absorbance of 0 at some wavelength means that no light of that particular wavelength has been absorbed. The intensities of the sample and reference beam are both the same, so the ratio (I_o/I) is 1 and the (log_<10>) of 1 is zero.

In a sample with an absorbance of 1 at a specific wavelength, what is the relative amount of light that was absorbed by the sample?

This question does not need Beer-Lambert Law (Equation ( ef<5>)) to solve, but only the definition of absorbance (Equation ( ef<2>))

[ A=log_ <10>left( dfrac ight) onumber]

The relative loss of intensity is

Equation ( ef<2>) can be rearranged using the properties of logarithms to solved for the relative loss of intensity:

Hence 90% of the light at that wavelength has been absorbed and that the transmitted intensity is 10% of the incident intensity. To confirm, substituting these values into Equation ( ef<2>) to get the absorbance back:


Complication

All assays have limits. Amounts of substance below some minimum will be undetectable. Beyond some maximum amount or concentration an assay becomes saturated, that is, increases in amount or concentration do not affect absorbance. We generally try to work within the linear range of an assay, that is, where absorbance is directly proportional to concentration. Ideally, we would set up standards that encompass the entire useful range of an assay. That is, we optimize the range of the assay.

Often a sample is so concentrated that when you assay the prescribed volume of sample the result is off scale &ndash the assay reagent is saturated. The solution then is to dilute the sample. For example, if the volume of each standard or sample is 1 ml, and 1 ml of your unknown gives a result that is off scale, you can add 0.1 ml sample to a test tube along with 0.9 ml buffer. If you read a concentration from the standard curve, then multiply the result by 10 to get the actual concentration in the sample. If you read an amount from the standard curve then simply divide that amount by 0.1 ml to get your concentration.

When samples are so concentrated that you cannot pipet a small enough amount accurately, you may have to conduct serial dilutions.


Conclusion:

In this experiment, a calibration curve was created by plotting absorbance vs. concentration in Excel. The calibration curve was constructed by measuring the absorbance rate of phosphate in five standard solutions.

The linear equation derived from the calibration curve was then manipulated and used to determine the concentration of phosphate in soda pop, and in an unknown water solution. The concentration of phosphate was experimentally determined to be 0.006834 M in Cola, and 1.41 x 10 -4 M in an unknown water sample.

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ANTICIPATED RESULTS

Typical results for the study of a globular protein (chicken hen’s egg white lysozyme) are given in Figure 2 and the results of the analysis of the spectra to give the secondary structure are given in Table 4 . The data collection of the 10 individual spectra use to create Figure 2 , including washing the cells between measurements, smoothing the data and correcting the data for the baseline and protein concentration took

4 hours and the 30 analyses summarized in Table 4 took

12 hours, including converting the data to the appropriate formats, running the calculations and tabulating the results.

Table 4

Analyses of the Structure of Lysozyme from the CD Data in Figure 2

Fraction of each secondary structure
MethodBasis SetsPath Length Cmconc. mg/mlwavelength range, nmhelix regularhelix endshelix totalbeta regularbeta endsbeta totalturnsother
CDPro Package a
X-Ray b 0.200.220.420.020.050.070.300.22
CDSSTR300.010.41260�0.190.140.330.110.080.190.200.28
CDSSTR300.010.83260�0.190.140.320.110.080.180.210.28
CDSSTR300.10.085260�0.200.130.340.140.080.220.190.25
CDSSTR300.10.41260�0.200.140.340.120.070.200.200.25
CONTIN300.010.41260�0.180.170.350.180.170.350.180.17
CONTIN300.010.83260�0.180.170.350.070.070.140.270.23
CONTIN300.10.085260�0.160.160.320.100.080.180.250.24
CONTIN300.10.41260�0.210.210.420.000.040.040.270.26
CONTINLL300.010.41260�0.190.160.340.080.070.150.230.27
CONTINLL300.010.83260�0.190.170.360.070.060.130.240.27
CONTINLL300.10.085260�0.170.150.320.110.080.180.240.25
CONTINLL300.10.41260�0.180.150.330.120.070.190.220.25
SELCON3300.010.41260�0.170.140.300.110.080.190.230.28
SELCON3300.010.83260�0.160.140.300.120.080.200.230.27
SELCON3300.10.085260�0.160.130.290.110.080.180.230.28
SELCON3300.10.41260�0.170.140.310.090.070.160.240.27
AVERAGE 0.180.160.340.100.080.170.230.26
STDEV 0.020.020.040.040.030.070.030.03
Other Methods
total helix totalbetaturnsother
X-RAY c 0.39 0.110.340.16
Method of constrained least squares fits with peptide references d
Peptide set 140.010.41240� 0.26 0.290.020.42
Peptide set 140.010.41240� 0.31 0.260.020.40
Peptide set 250.010.41240� 0.24 0.310.060.38
Peptide set 330.010.41240� 0.28 0.15NDND
Average 0.27 0.250.030.40
Stdev 0.03 0.070.020.02
Method of constrained least squares with references extracted from sets of protein spectra e
Protein Set 140.010.41240� 0.30 0.220.220.26
Protein Set 250.010.41240� 0.27 0.220.320.20
Methods that use singular value decomposition to deconvolute the reference data sets f
SVD330.010.41260� 0.27 0.150.170.25
VARSLC330.010.41260� 0.28 0.180.180.27
SELCON170.010.41260� 0.31 0.180.280.24
SELCON2170.010.41260� 0.32 0.160.240.29
Ridge Regression g
CONTIN g 160.010.41240� 0.30 0.300.230.18
Methods using neural network analyses h .
K2D180.010.41240� 0.33 0.14NDND
CDNN170.010.41260� 0.32 0.200.180.27
CDNN170.010.41260� 0.32 0.180.170.31
Average off all the methods using protein references excluding those in the CDPro a package
Average 0.30 0.190.220.25
Stdev 0.02 0.050.050.04

All of the methods used to analyze protein spectra should give reasonable estimates of α-helical content ( Table 4 ). The four programs included in the CDPro package, SELCON3, CONTIN, CONTINLL and CDSSTR should give results which are almost identical to each other and relatively independent of the lower limit wavelength range from 200 to 178 nm ( Table 4 ). Similar secondary contents should be also obtained with SELCON and SELCON2, CDNN and K2D ( Table 4 ). When proteins are analyzed using the method of least squares, basis sets extracted from proteins should give good fits, but some peptide data sets tend to over estimate β-structure and underestimate turns if data lower than 208 nm are fit 5 .

Some of the analysis programs output the fitted curve to the raw data. It should be noted that the programs giving the best fits to the data do not necessarily give the best estimates of protein conformation, because better fits will be obtained more variables are used. Therefore the fits obtained using CONTIN, which fits protein CD data by a large number of reference spectra, almost always gives a perfect fit to the raw data compared to methods that use fewer reference sets. Representative fits using programs with graphical output are shown in Figures 2e and 2f .


Thiol Redox Transitions in Cell Signaling, Part B: Cellular Localization and Signaling

Isaac K. Sundar , . Irfan Rahman , in Methods in Enzymology , 2010

7 HDACs Levels by Immunoblotting

In addition to HDAC activity, the levels of HDACs are also regulated by redox pathway via modulating their nucleocytoplasmic shuttling and posttranslational modifications, such as proteasome-dependent degradation ( Adenuga et al., 2009 Rahman et al., 2002 ). Therefore, protein level of HDACs is an important marker of oxidant-induced cell dysfunction or diseases such as COPD ( Ito et al., 2005 ).

In general, different cells fractions, such as whole lysates, and both cytoplasmic and nuclear protein are used to investigate the degradation and shuttling of HDACs ( Adenuga et al., 2009 Yang et al., 2006 ).

Protein estimation is done by the BCA, Bradford, or Lowry protein assay , following the manufacturer's instructions.

For HDAC1, HDAC2, HDAC3, SIRT1, and SIRT2 assays, 20 μg of isolated soluble proteins are electrophoresed on 7.5% SDS–PAGE gels, transferred onto nitrocellulose membranes (Amersham), and blocked with 10% nonfat dry milk in Tris-buffered saline (TBS) with 0.1% Tween-20 at 4 °C overnight.

Membranes are incubated with goat polyclonal anti-human anti-HDAC or anti-SIRT1 (1:1000 dilution in 5% nonfat dry milk in TBS) antibodies (HDAC1, SC-6298 HDAC2, SC-6296 HDAC3, SC-8138 from Santa Cruz Biotechnology SIRT1, Ab7343 SIRT2, Ab10659 from Abcam).

After washing, the levels of HDAC proteins are detected with rabbit anti-goat or anti-mouse antibody (1:20,000 dilution in 2.5% nonfat dry milk in TBS for 1 h) linked to horseradish peroxidase (Dako, Santa Barbara, CA), and bound complexes are detected with ECL (Jackson Immunology Research, West Grove, PA).


Enzyme Modification and Conjugation

1.1 Horseradish Peroxidase (HRP)

HRP (donor:hydrogen peroxide oxidoreductase EC 1.11.1.7), derived from horseradish roots, is a enzyme of molecular weight 40,000 that can catalyze the reaction of hydrogen peroxide with certain organic, electron-donating substrates to yield highly colored products ( Figure 22.1 ). The reaction of HRP with its fundamental substrate, H2O2, forms a stable intermediate that can dissociate in the presence of a suitable electron donor, oxidizing the donor and potentially creating a color change. The donor can consist of oxidizable molecules like ascorbate, cytochrome c, ferrocyanide, or the leuco forms of many dyes. A large variety of electron-donating dye substrates are commercially available for use as HRP detection reagents. Some of them can be used to form soluble colored products for use in spectrophotometric detection systems, while other substrates form insoluble products that are especially appropriate for staining techniques. In addition, substrates are available that create fluorescent or chemiluminescent products upon oxidation with HRP. The chemiluminescent substrates are among the most sensitive of all detection reagents, facilitating the detection of as little as attogram quantities of many targeted analytes. The pH optimum for HRP is 7.0 although particular substrate detection reactions may be performed at pH values slightly different from neutrality.

Figure 22.1 . Horseradish peroxidase shown as the ribbon structure with the heme ring in its active center and two bound calcium ions. The molecular model is based on structure 1H58 in the RCSB Protein Databank by Berglund et al. (2002) .

The use of antibody–HRP or streptavidin–HRP conjugates in peroxidase-catalyzed enhanced chemiluminescent assays can result in one of the most sensitive detection methods for assaying targeted analytes in ELISA and western blotting applications. The reaction cascade that occurs during HRP catalysis can be dramatically improved by the addition of various enhancer molecules, which create oxidized intermediates leading to the oxidation and light emission of a chemiluminescent substrate such as luminol. Vdovenko et al. (2012) analyzed this reaction using a multi-factorial design of experiments (DOE) approach to identify the best combination and concentrations of H2O2, luminol, and two different enhancer compounds (3-(10′-phenothiazinyl)propane-1-sulfonate and 4-morpholinopyridine. This combination at an optimized concentration resulted in the best signal-to-noise ratio and the longest chemiluminescent emission.

HRP is a hemoprotein containing photohemin IX as its prosthetic group. The presence of the heme structure gives the enzyme its characteristic color and maximal absorptivity at 403 nm. The ratio of its absorbance in solution at 403 nm to its absorbance at 275 nm, called the RZ or Reinheitzahl ratio, can be used to approximate the purity of the enzyme. However, at least seven isoenzymes exist for HRP ( Shannon et al., 1966 Kay et al., 1967 Strickland et al., 1968 ), and their RZ values vary from 2.50 to 4.19. Thus, unless the RZ ratio is precisely known or determined for the particular isoenzyme of HRP utilized in the preparation of an antibody–enzyme conjugate, subsequent measurement after crosslinking would yield questionable results in the determination of the amount of HRP present in the conjugate.

HRP is a glycoprotein that contains significant amounts of carbohydrate. Its polysaccharide chains are often used in crosslinking reactions to couple the enzyme to targeting molecules. Mild oxidation of its associated glycan sugar residues with sodium periodate generates reactive aldehyde groups that can be used for conjugation to amine-containing molecules. Reductive amination of oxidized HRP to antibody molecules in the presence of sodium cyanoborohydride is perhaps the simplest method of preparing highly active conjugates with this enzyme ( Chapter 4, Section 1.4 , and Chapter 20, Section 1.3 ).

Other methods of HRP conjugation include the use of the homobifunctional reagent glutaraldehyde ( Chapter 5, Section 6.2 , and Chapter 15, Section 2.1 ) and the heterobifunctional crosslinker, SMCC (succinimidyl-4-(N-maleimidomethyl)cyclohexane-1-carboxylate) ( Chapter 6, Section 1.3 ). Using glutaraldehyde, a two-step protocol is usually employed to try to limit the extent of oligomer formation. Even using the most highly controlled reactions, however, this method often causes unacceptable amounts of precipitated conjugate. Despite this disadvantage, glutaraldehyde conjugation is still routinely used, especially in the preparation of some antibody–enzyme reagents that go into established diagnostic assays. The use of the N-hydroxysuccinimide (NHS) ester–maleimide crosslinker, SMCC, provides far better control over the conjugation process. SMCC is usually reacted first with HRP to create a derivative containing sulfhydryl-reactive maleimide groups. HRP activation of the native enzyme should result in the modification of a maximum of about two amine groups on the protein, because HRP only contains two lysines. An increase in the activation level can be realized if the enzyme first is modified with ethylenediamine (EDA) using the carbodiimide EDC according to the methods described in Chapter 19 for the production of cationized bovine serum albumin (cBSA). The EDA-modified HRP is also more stable than the unmodified version, so cationization may have benefits in the retention of enzyme activity. The maleimide-activated enzyme can be purified and freeze-dried, providing a ready source of modified HRP to react with a sulfhydryl-containing antibody. Several preactivated forms of this enzyme are available from Thermo Fisher.

The size of HRP is an advantage in preparing antibody–enzyme conjugates, since the overall complex size also can be designed to be small. Relatively low-molecular-weight conjugates are able to penetrate cellular structures better than large, polymeric complexes. This is why HRP conjugates are often the best choice for immunohistochemical (IHC) and immunocytochemical staining techniques. Small conjugate size means greater accessibility to antigenic structures within tissue sections.

Another distinctive advantage of HRP is its robust nature and stability, especially under the conditions employed for crosslinking. HRP is stable for years in a freeze-dried state, and the purified enzyme can be stored in solution at 4°C for many months without significant loss of activity. The enzyme also retains excellent activity after being modified with a conjugation reagent or after being periodate-oxidized to form aldehyde groups on its polysaccharide chains. Depending on the methods used for crosslinking, HRP conjugates can be constructed to have a high ratio of enzyme to antibody or a low ratio—both retaining high specific activity.

The disadvantages associated with HRP are several. The enzyme only contains two available primary ε-amine groups—extraordinarily low for most proteins—thus limiting its ability to be activated with amine-reactive heterobifunctionals. HRP is sensitive to the presence of many antibacterial agents, especially azide. It is also reversibly inhibited by cyanide and sulfide ( Theorell, 1951 ). Finally, while the enzymatic activity of HRP is extremely high, its useful life span or practical substrate development time is somewhat limited. After about an hour of substrate turnover, in some situations its activity can be decreased severely.

Nevertheless, HRP is by far the most popular enzyme used in antibody–enzyme conjugates. One survey of enzyme use stated that HRP is incorporated in about 80% of all antibody conjugates, most of them utilized in diagnostic assay systems.


Ultraviolet-Visible (UV-Vis) Spectroscopy

Ultraviolet-visible (UV-Vis) spectroscopy is one of the most popular analytical techniques because it is very versatile and able to detect nearly every molecule. With UV-Vis spectroscopy, the UV-Vis light is passed through a sample and the transmittance of light by a sample is measured. From the transmittance (T), the absorbance can be calculated as A=-log (T). An absorbance spectrum is obtained that shows the absorbance of a compound at different wavelengths. The amount of absorbance at any wavelength is due to the chemical structure of the molecule.

UV-Vis can be used in a qualitative manner, to identify functional groups or confirm the identity of a compound by matching the absorbance spectrum. It can also be used in a quantitative manner, as concentration of the analyte is related to the absorbance using Beer's Law. UV-Vis spectroscopy is used to quantify the amount of DNA or protein in a sample, for water analysis, and as a detector for many types of chromatography. Kinetics of chemical reactions are also measured with UV-Vis spectroscopy by taking repeated UV-Vis measurements over time. UV-Vis measurements are generally taken with a spectrophotometer. UV-Vis is also a very popular detector for other analytical techniques, such as chromatography, because it can detect many compounds.

Typically, UV-Vis is not the most sensitive spectroscopy technique, because not a lot of light is absorbed over a short path length. Other spectroscopy techniques such as fluorescence have higher sensitivity, but they are not as generally applicable, as most molecules are not fluorescent. UV-Vis has a similar sensitivity to other absorbance measurements, such as infrared spectroscopy.

Principles

UV-Vis is often called a general technique because most molecules will absorb in the UV-Vis wavelength range. The UV extends from 100� nm and the visible spectrum from 400� nm. The 100� nm range is called the deep UV. Light sources are more difficult to find for this range, so it is not routinely used for UV-Vis measurements. Typical UV-Vis spectrometers use a deuterium lamp for the UV that produces light from 170� nm and a tungsten filament lamp for visible, which produces light from 350𔃀,500 nm.

When a photon hits a molecule and is absorbed, the molecule is promoted into a more excited energetic state. UV-visible light has enough energy to promote electrons to a higher electronic state, from the highest occupied molecular orbital (HOMO) to the lowest unoccupied molecular orbital (LUMO). The energy difference between the HOMO and the LUMO is called the band gap. Typically, these orbitals are called bonding and anti-bonding. The energy of the photon must exactly match the band gap for the photon to be absorbed. Thus, molecules with different chemical structures have different energy band gaps and different absorption spectra. The most common transitions that fall in the UV-Vis range are π-π* and n- π*. Pi orbitals arise due to double bonds, and n orbitals are for non-bonding electrons. Pi star are anti-bonding pi orbitals. Thus, the best UV-Vis absorption is by molecules that contain double bonds. Pi orbitals adjacent to each other that are connected, called conjugation, typically increases absorption. Sigma-σ* transitions, associated with single bonds, are higher energy and fall in the deep UV, so they are less useful for routine use. The appearance of broad bands or shoulders on the UV-Vis structure is due to the numerous vibrational and rotational states of a molecule, which lead to separate energy band gaps of slightly different energies.

For molecules with absorption in the visible region, the compounds will often appear colored. However, a common misconception is that the wavelength of peak absorption (λmax) for a compound is the color it appears. A compound that appears red does not have much absorption in the red region of the spectrum. Instead, the λmax for a compound that looks red is green. The color of a compound arises because those wavelengths of light are selectively transmitted through the sample, and thus they are not absorbed. A color wheel is helpful in determining what color a compound will absorb and what range the λmax will be, as the color directly across the wheel from the observed color is the color that is most absorbed.

Absorption follows Beer's Law, A=εbC where ε is the molar attenuation coefficient, b is path length, and C is concentration. The molar attenuation coefficient is the characteristic of an individual compound to absorb at a given wavelength and this property is due to functional groups, conjugation, etc. If a compound does not have a high attenuation coefficient , it could be tagged with an appropriate group to increase its absorbance. Path length is generally related to the size of the cuvette and is 1 cm in standard spectrophotometers.

UV-Vis is performed on a variety of instruments, from traditional spectrophotometers to more modern-day plate readers. The absorbance wavelength must be chosen, either using a filter or a monochromator. A monochromator is a device that separates the wavelengths of light spatially and then places an exit slit where the desired wavelength of light is. Monochromators can be scanned to provide a whole absorbance spectrum. Alternatively, a diode-array instrument allows all colors of light to be transmitted through the sample, then the light is separated into different wavelengths spatially and detected using photodiodes. Diode-array instruments collect full spectra faster, but are more complicated and more expensive.

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Procedure

1. Calibrate the Spectrometer

  1. Turn on the UV-Vis spectrometer and allow the lamps to warm up for an appropriate period of time (around 20 min) to stabilize them.
  2. Fill a cuvette with the solvent for the sample and make sure the outside is clean. This will serve as a blank and help account for light losses due to scattering or absorption by the solvent.
  3. Place the cuvette in the spectrometer. Make sure to align the cuvette properly, as often the cuvette has two sides, which are meant for handling (may be grooved) and are not meant to shine light through.
  4. Take a reading for the blank. The absorbance should be minimal, but any absorbance should be subtracted out from future samples. Some instruments might store the blank data and perform the subtraction automatically.

2. Perform an Absorbance Spectrum

  1. Fill the cuvette with the sample. To make sure the transfer is quantitative, rinse the cuvette twice with the sample and then fill it about ¾ full. Make sure the outside is clean of any fingerprints, etc.
  2. Place the cuvette in the spectrometer in the correct direction.
  3. Cover the cuvette to prevent any ambient light.
  4. Collect an absorbance spectrum by allowing the instrument to scan through different wavelengths and collect the absorbance. The wavelength range can be set with information about the specific sample, but a range of 200� nm is standard. A diode-array instrument is able to collect an entire absorbance spectrum in one run.
  5. From the collected absorbance spectrum, determine the absorbance maximum (λmax). Repeat the collection of spectra to get an estimate of error in λmax.
  6. To make a calibration curve, collect the UV-Vis spectrum of a variety of different concentration samples. Spectrometers are often limited in linear range and will not be able to measure an absorbance value greater than 1.5. If the absorbance values for the sample are outside the instrument's linear range, dilute the sample to get the values within the linear range.

3. Kinetics Experiments with UV-Vis Spectroscopy

  1. UV-Vis can be used for kinetics experiments by examining the change in absorbance over time. For a kinetics experiment, take an initial reading of the sample.
  2. Quickly add the reagent to start the chemical reaction.
  3. Stir it well to mix with the sample. If a small amount is added, this could be done in a cuvette. Alternatively, mix the reagent with sample and quickly pour some in a cuvette for a measurement.
  4. Measure the absorbance at the λmax for the analyte of interest over time. If using up the reagent being measuring (i.e. absorbance is going up because there is less reagent to absorb), then the decay will indicate the order of the reaction.
  5. Using a calibration curve, make a plot of analyte concentration vs time, converting the absorbance value into concentration. From there, this graph can be fit with appropriate equations to determine the reaction rate constants.

Ultraviolet-visible, or UV-Vis, spectroscopy is one of the most popular analytical techniques in the laboratory.

In UV-Vis spectroscopy, light is passed through a sample at a specific wavelength in the UV or visible spectrum. If the sample absorbs some of the light, not all of the light will be pass through, or be transmitted. Transmission is the ratio of the intensity of the transmitted light to the incident light, and is correlated to absorbance. The absorbance can be used in a quantitative manner, to obtain the concentration of a sample. It can also be used in a qualitative manner, to identify a compound by matching the measured absorbance over a range of wavelengths, called the absorbance spectrum, to the published data. This video will introduce UV-Vis spectroscopy, and demonstrate its use in the laboratory in determining sample concentration and reaction kinetics.

When a photon hits a molecule and is absorbed, the molecule is promoted from its ground state into a higher energy state. The energy difference between the two is the band gap. The energy of the photon must exactly match the band gap in order for the photon to be absorbed. The chemical structure determines the band gap therefore molecules each have unique absorbance spectra.

Absorbance follows Beer's Law, which states absorbance equals the molar attenuation coefficient times the path length and concentration. The molar attenuation coefficient is related to the individual compound's ability to absorb light of a specific wavelength. Path length refers to the distance traveled by light through the sample, which is typically 1 cm for standard cuvettes. Beer's law can be used to calculate sample concentration, if the absorptivity is known, or a calibration curve can be used.

UV-Vis is often called a general technique, as most molecules absorb light in the UV-visible wavelength range. The UV range extends from 100� nm, and the visible spectrum ranges from 400� nm. However, most spectrophotometers do not operate in the deep UV range of 100� nm, as light sources in this range are expensive. Most UV-Vis spectrophotometers use a deuterium lamp for the UV range, which produces light from 170� nm, and a tungsten filament lamp for the visible range, which produces light from 350𔃀,500 nm.

Since the light source is usually a lamp with broad wavelength ranges, the specific absorbance wavelength is selected using filters or a monochromator. A monochromator is a device that separates the wavelengths of light spatially, and then places an exit slit where the desired wavelength of light is. The monochromator can be scanned over a wavelength range to provide an entire absorbance spectrum. This makes the technique useful for quantifying and identifying a wide range of molecules.

Now that the basics of UV-Vis spectroscopy have been outlined, lets take a look at a simple UV-Vis experiment in the laboratory.

Before beginning the measurement, turn on the spectrophotometer, and allow the lamps to warm up for an appropriate period of time to stabilize them.

Prepare a blank by filling a clean cuvette with the sample solvent, and then wipe the outside with lint-free paper to remove any fingerprints.

Ensure that the cuvette is aligned properly with any grooved sides out of the beam-path, and insert it into the spectrophotometer. Secure the lid to prevent ambient light from entering the system.

Measure the absorbance of the blank at one wavelength, or over a wavelength range. Record or save the absorbance, as it must be subtracted from the absorbance of the sample.

Next, discard the blank and rinse the cuvette twice with sample. Then, fill the cuvette about ¾ full with sample. Wipe the outside of the cuvette again, to ensure that it is clean and free of fingerprints.

Place the cuvette in the spectrophotometer in the correct orientation, and secure the lid.

Collect an absorbance measurement or spectrum at the same wavelength or wavelength range as the blank. Subtract the blank spectrum or measurement, if the instrument does not automatically do so.

From the collected absorbance spectrum, determine the absorbance maximum, or λmax.

To quantify the amount of analyte in the sample, create a calibration curve using a range of known analyte concentrations. For more information on how to construct and use a calibration curve, please watch this collection's video "Calibration Curves".

The absorbance measurement can also be used to calculate reaction kinetics by measuring the increase or decrease in a compounds concentration throughout the reaction. Begin by taking an initial reading of the sample, blue dye in this case, at the absorbance maximum before the reaction.

Next, quickly add the reagent, bleach in this case, to start the chemical reaction. Stir it well, so that it mixes with the sample.

Measure the absorbance at the absorbance maximum over time.

The initial absorbance spectrum of the blue dye sample is shown. The background colors show the colors of light in the visible spectrum. The blue dye has an absorbance maximum at about 630 nm.

The kinetics of the reaction between blue dye and bleach was measured over time. The absorbance of blue dye decreases over time, as it reacts with the bleach. The absorbance reaches near zero after 300 s, indicating that the reaction has neared completion. For more information on kinetics and reactions, please watch the JoVE Science Education video "Reaction Rate Laws".

UV-Vis spectroscopy is used heavily in many different research areas to identify or quantify a sample.

For example, UV-Vis spectroscopy is used heavily in biological fields to quantify the amount of protein in a sample. A Bradford assay is often used to quantify proteins, with the aid of a dye. First, a calibration curve of known protein concentrations is prepared, typically using Bovine Serum Albumin, or BSA. Then Coomassie blue stain is added to each of the standards and to the sample. The absorbance of the protein-dye complex is then measured at 595 nm.

Alternatively, proteins can be measured directly by their absorbance at 280 nm. In this example, protein concentration is quantified using an ultra low volume spectrophotometer. For many proteins, an absorbance of 1 correlates to a concentration of 1 mg/mL.

UV-Vis spectroscopy is also used to quantify the amount of bacterial cells in a cell culture. For this measurement, the absorbance, or optical density, is measured at 600 nm. Typically, an OD600 measurement of 1 indicates the presence of 8 x 10 8 bacterial cells per mL. Measuring the cell density throughout culture growth enables the determination of the bacterial growth curve, and can help to identify when a culture is in its exponential growth phase.

Nitrogen oxide and nitrogen dioxide, or NOx, is a by-product of automobile exhaust, and can be harmful to the environment because it forms damaging tropospheric ozone. NOx can be measured by reacting it with a solution of sulfanilic acid and napthyl-ethylenediamine. The resulting solution is a pink colored azo dye molecule, the intensity of which is directly correlated to NOx concentration. This concentration can then be determined using a UV-Vis spectrophotometer.

You've just watched JoVE's introduction to UV-visible spectroscopy. You should now understand the basics of UV-Vis operation, how to measure a sample using a UV-Vis and how to correlate absorbance to sample concentration.

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Results

UV-Vis can be used to obtain a spectrum of colored compounds. In Figure 1A, the absorbance spectrum of a blue dye is shown. The background shows the colors of light in the visible spectrum. The blue dye has a λmax absorbance in the orange/red. Figure 1B shows a spectrum of a red dye, with λmax in the green.

Kinetics can be measured from a plot of absorbance at one wavelength over time. Figure 2 shows a plot of the absorbance of a blue dye (at 630 nm) as it reacts with bleach.


Figure 1. UV-Vis absorbance spectra. A. Blue dye #1 has maximum absorbance in the orange/red. B. Red dye #40 has maximum absorbance in the green. Please click here to view a larger version of this figure.


Figure 2. UV-Vis for kinetics. Absorbance of blue dye #1 as it reacts with bleach. The curve can be fit with an exponential decay, indicating first order kinetics. Please click here to view a larger version of this figure.

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Applications and Summary

UV-Vis is used in many chemical analyses. It is used to quantitate the amount of protein in a solution, as most proteins absorb strongly at 280 nm. Figure 3 shows an example spectra of cytochrome C, which has a high absorbance at 280 and also at 450 because of a heme group. UV-Vis is also used as a standard technique to quantify the amount of DNA in a sample, as all the bases absorb strongly at 260 nm. RNA and proteins also absorb at 260 nm, so absorbance at other wavelengths can be measured to check for interferences. Specifically, proteins absorb strongly at 280 nm, so the ratio of absorbance at 280/260 can give a measure of the ratio of protein to DNA in a sample.

Most simple analyses measure the absorbance one wavelength at a time. However, more chemical information is present if measurements are made at many wavelengths simultaneously. Diode-array instruments capture all the light that is transmitted, split the light into different colors using a prism or holographic grating, and then absorbance at different wavelengths is captured on a linear array of photodiodes. The advantage of this method is that it is useful for measuring many different molecules simultaneously.


Figure 3. UV-Vis spectrum of a protein. The peak at 280 nm is indicative of a protein. The peak at 450 is due to absorbance of the heme group in cytochrome C.

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Transcript

Ultraviolet-visible, or UV-Vis, spectroscopy is one of the most popular analytical techniques in the laboratory.

In UV-Vis spectroscopy, light is passed through a sample at a specific wavelength in the UV or visible spectrum. If the sample absorbs some of the light, not all of the light will be pass through, or be transmitted. Transmission is the ratio of the intensity of the transmitted light to the incident light, and is correlated to absorbance. The absorbance can be used in a quantitative manner, to obtain the concentration of a sample. It can also be used in a qualitative manner, to identify a compound by matching the measured absorbance over a range of wavelengths, called the absorbance spectrum, to the published data. This video will introduce UV-Vis spectroscopy, and demonstrate its use in the laboratory in determining sample concentration and reaction kinetics.

When a photon hits a molecule and is absorbed, the molecule is promoted from its ground state into a higher energy state. The energy difference between the two is the band gap. The energy of the photon must exactly match the band gap in order for the photon to be absorbed. The chemical structure determines the band gap therefore molecules each have unique absorbance spectra.

Absorbance follows Beer's Law, which states absorbance equals the molar attenuation coefficient times the path length and concentration. The molar attenuation coefficient is related to the individual compound's ability to absorb light of a specific wavelength. Path length refers to the distance traveled by light through the sample, which is typically 1 cm for standard cuvettes. Beer's law can be used to calculate sample concentration, if the absorptivity is known, or a calibration curve can be used.

UV-Vis is often called a general technique, as most molecules absorb light in the UV-visible wavelength range. The UV range extends from 100–400 nm, and the visible spectrum ranges from 400–700 nm. However, most spectrophotometers do not operate in the deep UV range of 100–200 nm, as light sources in this range are expensive. Most UV-Vis spectrophotometers use a deuterium lamp for the UV range, which produces light from 170–375 nm, and a tungsten filament lamp for the visible range, which produces light from 350–2,500 nm.

Since the light source is usually a lamp with broad wavelength ranges, the specific absorbance wavelength is selected using filters or a monochromator. A monochromator is a device that separates the wavelengths of light spatially, and then places an exit slit where the desired wavelength of light is. The monochromator can be scanned over a wavelength range to provide an entire absorbance spectrum. This makes the technique useful for quantifying and identifying a wide range of molecules.

Now that the basics of UV-Vis spectroscopy have been outlined, lets take a look at a simple UV-Vis experiment in the laboratory.

Before beginning the measurement, turn on the spectrophotometer, and allow the lamps to warm up for an appropriate period of time to stabilize them.

Prepare a blank by filling a clean cuvette with the sample solvent, and then wipe the outside with lint-free paper to remove any fingerprints.

Ensure that the cuvette is aligned properly with any grooved sides out of the beam-path, and insert it into the spectrophotometer. Secure the lid to prevent ambient light from entering the system.

Measure the absorbance of the blank at one wavelength, or over a wavelength range. Record or save the absorbance, as it must be subtracted from the absorbance of the sample.

Next, discard the blank and rinse the cuvette twice with sample. Then, fill the cuvette about ¾ full with sample. Wipe the outside of the cuvette again, to ensure that it is clean and free of fingerprints.

Place the cuvette in the spectrophotometer in the correct orientation, and secure the lid.

Collect an absorbance measurement or spectrum at the same wavelength or wavelength range as the blank. Subtract the blank spectrum or measurement, if the instrument does not automatically do so.

From the collected absorbance spectrum, determine the absorbance maximum, or λmax.

To quantify the amount of analyte in the sample, create a calibration curve using a range of known analyte concentrations. For more information on how to construct and use a calibration curve, please watch this collection's video "Calibration Curves".

The absorbance measurement can also be used to calculate reaction kinetics by measuring the increase or decrease in a compounds concentration throughout the reaction. Begin by taking an initial reading of the sample, blue dye in this case, at the absorbance maximum before the reaction.

Next, quickly add the reagent, bleach in this case, to start the chemical reaction. Stir it well, so that it mixes with the sample.

Measure the absorbance at the absorbance maximum over time.

The initial absorbance spectrum of the blue dye sample is shown. The background colors show the colors of light in the visible spectrum. The blue dye has an absorbance maximum at about 630 nm.

The kinetics of the reaction between blue dye and bleach was measured over time. The absorbance of blue dye decreases over time, as it reacts with the bleach. The absorbance reaches near zero after 300 s, indicating that the reaction has neared completion. For more information on kinetics and reactions, please watch the JoVE Science Education video "Reaction Rate Laws".

UV-Vis spectroscopy is used heavily in many different research areas to identify or quantify a sample.

For example, UV-Vis spectroscopy is used heavily in biological fields to quantify the amount of protein in a sample. A Bradford assay is often used to quantify proteins, with the aid of a dye. First, a calibration curve of known protein concentrations is prepared, typically using Bovine Serum Albumin, or BSA. Then Coomassie blue stain is added to each of the standards and to the sample. The absorbance of the protein-dye complex is then measured at 595 nm.

Alternatively, proteins can be measured directly by their absorbance at 280 nm. In this example, protein concentration is quantified using an ultra low volume spectrophotometer. For many proteins, an absorbance of 1 correlates to a concentration of 1 mg/mL.

UV-Vis spectroscopy is also used to quantify the amount of bacterial cells in a cell culture. For this measurement, the absorbance, or optical density, is measured at 600 nm. Typically, an OD600 measurement of 1 indicates the presence of 8 x 108 bacterial cells per mL. Measuring the cell density throughout culture growth enables the determination of the bacterial growth curve, and can help to identify when a culture is in its exponential growth phase.

Nitrogen oxide and nitrogen dioxide, or NOx, is a by-product of automobile exhaust, and can be harmful to the environment because it forms damaging tropospheric ozone. NOx can be measured by reacting it with a solution of sulfanilic acid and napthyl-ethylenediamine. The resulting solution is a pink colored azo dye molecule, the intensity of which is directly correlated to NOx concentration. This concentration can then be determined using a UV-Vis spectrophotometer.

You've just watched JoVE's introduction to UV-visible spectroscopy. You should now understand the basics of UV-Vis operation, how to measure a sample using a UV-Vis and how to correlate absorbance to sample concentration.


Overview

VivoTags ® are optimized NIR labels for labeling molecules including peptides, small molecules, proteins, macromolecules, and nanoparticles. They are available as either NHS esters or maleimide reactive dyes which makes conjugation to either free amine (-Nh3) and free thiol (-SH) containing molecules possible. VivoTags have validated for both in vivo and in vitro applications. Figure 1 below shows the conjugation reaction.


Figure 1: Conjugation reaction for NHS ester (A) and maleimide (B) VivoTag dyes. VivoTag labeled conjugates have been validated in both IVIS ® and FMT ® imaging systems. In Figure 2 below, Nu/nu mice were orthotopically injected with the Bioluminescent cell line MDA-MB-231-luc2. An antibody-conjugate labeled with VivoTag with specifity for the tumor cells was injected and compared to a VivoTag labeled IgG with no tumor specificity.

Figure 2: Imaging VivoTag conjugates on IVIS and FMT systems. A. Bioluminescence (left) and fluorescence intensity (right) were measured using an IVIS spectrum. In both mice the biolumenscence from the tumor cell line is strong, but only the mouse on the right, injected with the tumor specific VivoTag conjugate shows fluorescence at the tumor site. B. FMT reflectance mode is used to show the mouse on the right, injected with the tumor specific VivoTag conjugate, s a strong signal in the area of the of the orthotopically injected tumor. The high signal in the liver is due the distribution kinetics of the antibody. All images shown are at 24 hours post injection of the antibody conjugate.

Products and catalog numbers

Choosing the right VivoTag labeling agent

  • Dye Wavelength: Be sure to choose the proper wavelength that your instrument is capable of measuring effectively. For in vitro applications such as microscopy, dyes above 750 nm can't typically be imaged. For deep tissue measurements in vivo however, longer wavelength dyes such as VivoTag 750 or VivoTag 800 are preferred provided your in vivo imager has the proper filter sets.
  • VivoTag vs. VivoTag MAL: VivoTags are suitable for labeling free amines (typically lysine residues), while VivoTag MAL dyes are suitable for labeling free thiols (typically cysteine residues).
  • VivoTag vs. VivoTag -S: The "S" in VivoTag -S stands for "self-quenching". When a protein is labeled with multiple VivoTag-S molecules, there is a shift in the absorbance of the dye, which will ultimately reduce the signal if using the filter sets listed above. Only choose VivoTag -S if you are sure your labeling ratio of dye: protein will be 1:1. If multiple dyes are to be conjugated onto your protein, the non -"S" version should be chosen.
  • Package Size: Standalone Vivotags are available in 1 mg and 5 mg package sizes. Typically you can label 10-20 mg of antibody using 1 mg of dye, but we do recommend trying different dye: protein conjugation ratios to determine the optimal conditions for your protein of interest.

What do I need to perform a VivoTag conjugation?

  • Appropriate VivoTag labeling reagent (see Table 1 in section above for catalog numbers)
  • Appropriate conjugation buffer
    • For NHS ester, 50 mM carbonate/biocarbonate buffer, pH 8.3-8.5
    • For maleimide, appropriate buffer (e.g. 100 mM PBS, 100 mM TRIS, or 100 mM HEPES buffer, pH 6.5-7.0)
    • For NHS ester, Bio-Rad Bio-Gel P-100
    • For Maleimide, Sephadex G-25
    • 0.2 µm syringe filter to filter the purified conjugate
    • Absorbance reader (UV spectrophotometer, NanoDrop spectrophotometer) to determine the degree of labeling

    Protocols and calculations

    General overview of conjugation protocol

    Determining the degree of labeling

    In order to determine the degree of labeling, the absorbance of the diluted VivoTag-protein conjugate is measured at both 280 nm (to determine the amount of protein present) and at the absorbance wavelength of the VivoTag dye (643 nm to 790 nm, depending on the dye-See Table 1 above). There is a small amount of crosstalk at 280 nm as the VivoTag dyes also absorb at this wavelength, so it is necessary to subtract out this background absorbance in order to precisely calculate the degree of labeling. The amount of crosstalk varies by which VivoTag dye is chosen. Table 2 lists the amount of crosstalk for each VivoTag at 280 nm.

    VivoTag% Crosstalk at 280 nmCorrection factor (Cf)
    VivoTag 64550.05
    VivoTag 645 MAL50.05
    VivoTag 680 XL160.16
    VivoTag 680 XL MAL130.13
    VivoTag-S 680160.16
    VivoTag-S 75060.06
    VivoTag-S 750 MAL50.05
    VivoTag 80050.05
    Table 2: Correction factors for VivoTags. The correction factor is used to calculate the background absorbance of the dye at 280 nm. See sample calculation below.

    Sample Calculation
    Here is an example of the calculation if you labeled of an IgG labeled with VivoTag 645. After purification, you dilute the conjugate 1:10 in PBS. You measure the A280 of the dilution to be 0.415 and the A643 of the dilution to be 0.786. The Epsilon (Ɛ) for IgG is 210,000 M -1 cm -1 and the Epsilon (Ɛ) for VivoTag 645 is 210,000 M -1 cm -1 (see Table 1 for all dye Epsilons).
    First the A280 and the A643 need to be multiplied by the dilution factor of 10 in order to determine the stock concentration:

    A280 = 0.415 x 10 = 4.15
    A643 = 0.786 x 10 = 7.86

    To calculate the concentration of the IgG (Pc) the correction factor needs to be multiplied by the dye absorbance and subtracted from the A280:

    Pc [M] = (A280 - (A643)*Cf)/Ɛ of IgG
    Pc = (4.15 - 7.86*0.05)/210,000
    Pc = 4.15-0.393/210,000
    Pc = 3.757/210,000 = .00001789 M or 17.89 µM IgG

    To calculate the concentration of the VivoTag 645 dye:

    Dc [M] = (A643)/Ɛ of VivoTag 645
    Dc = 7.86/210,000
    Dc = 0.0000374 M or 37.4 µM VivoTag 645

    To calculate the degree of labeling:

    In this case, there are 2.09 VivoTag dye molecules for every molecule of IgG. Ideal dye-to-protein labeling ratios are typically between 2 and 3. It is important to note that the above equations assume a path length of 1 cm (typical path length of a cuvette). If you are using a NanoDrop spectrophotometer to measure absorbance, the path length is only 1 mm so A280 and A643 values obtained by this method would have to multiplied by an additional factor of 10. Omitting this correction factor will not alter the dye-to-protein ratio calculated, but will alter the µM of dye measured which will affect the amount of conjugate injected into your in vivo model organism.

    Assay optimization in vivo

    Not all proteins or antibodies make good imaging agents, independent of target specificity. Some may have too long or short of a half-life in vivo, or may excessively accumulate in non-target sites. It is a good idea to test specificity both in vitro and in vivo is possible, and to determine the in vivo biodistribution and pharmacokinetics of your VivoTag conjugate.

    General Product and Application FAQs

    Q. What is the structure of VivoTag?
    A. Unfortunately the structure of VivoTag is proprietary and we are unable to disclose it to customers.

    Q. What is the difference between VivoTag and VivoTag -S?
    A. VivoTag-S refers to a shift in absorbance that the dye experiences when conjugated on a molecule with multiple dyes. The shift in absorbance upon conjugation is about 50-60 nm lower in wavelength than the free dye alone.

    Q. For VivoTag -S what does the shift in absorbance correspond to in terms of what I'll see on my instrument? Do I need to adjust any filter settings?
    A. It depends on the instrument that is being used. If multiple tags in close proximity are added then the new max wavelength will be 50-60 nm toward the blue range and if it is only one VivoTag the wavelength will be the one reported on the technical data sheet.

    Q. Will there be an increase or decrease in signal based on the number of VivoTag molecules that bind to my protein?
    A. This will depend on the protein being used and the method of detection being used as well.

    Q. What is the difference between VivoTag and VivoTag MAL?
    A. The main difference between the regular dyes and the MAL dyes is that the MAL dyes are better suited for labeling thiol and cysteine containing molecules. The dye properties (abs, FL, ex. coeff, etc) are all the same.

    Q. Can VivoTags be used to label cells?
    A. While this is possible, we instead recommend VivoTrack™ for this application.

    Q. Can I use VivoTags in humans?
    A. No. VivoTags are intended for research purposes only and are not for human use.

    Q. When optimizing my experiment in vivo, is there a control dye that can be used?
    A. Yes, you can use Genhance™ as a control to determine non-specific uptake as this dye has no specificity for any particular target in vivo (other than vasculature). It is the acid form of the VivoTag dyes and is available as in 680 nm and 750 nm versions. Alternatively, you could conjugate a non-targeting protein or antibody to VivoTag and measure its specificity compared to your targeted molecule.

    Labeling and Purification FAQs

    Q. Can I reconstitute a VivoTag in an aqueous buffer?
    A. Yes you technically can if your experiment requires it, but the reaction needs to be done very quickly so that the NHS group does not hydrolyze and render the dye no longer reactive.

    Q. What amount of dye is a good starting point for protein or antibody labeling?
    A. Typically you can label 10-20 mg of antibody or protein with 1 mg of dye. This is a good starting point, but since all molecules are unique the amounts and ratios need to be optimized.

    Q. I do not want to run my conjugate over a column to purify. Can I dialyze away the free dye instead?
    A. For the NHS labeling reagents, we not recommend dialysis as a way to remove free dye. For the maleimide dyes, dialysis can be used instead of a gel filtration column if desired.

    Q.What mass should I use when analyzing my NHS VivoTag conjugates using mass spec?
    A. This table contains the mass expected when using mass spec analysis:

    VivoTagMass (X) to be added to the MW of biomolecule
    VivoTag 645974.2
    VivoTag 680 XL1234.2
    VivoTag-S 6801019.2
    VivoTag-S 750865.2
    VivoTag 8001045.2

    Q. Why does the table above contain values for the molecular weight that differ from the technical data sheet?
    A.The VivoTags exist as triethylammonium series salts. The number of triethylammonium groups vary from bis triethylammonium to penta triethyammonium dyes. When running Mass Spec, depending on the conditions, it is possible to observe up to 5*101(MW of TEA) less than the number reported on the TD sheet. Also upon Mass Spec analyses the dye will hydrolyze giving the MW of the acid form rather than the NHSE form of the dye.

    Storage and Stability FAQs

    Q.How long is the VivoTag powder good for?
    A. Stability of the powder will vary for each product. Please check the technical data sheet for the most accurate information about storage temperature and shelf life.

    Q. How long is the reconstituted VivoTag dye good for?
    A. We recommend storing the DMSO reconstituted VivoTag dye at 4 °C protected from light. Under these conditions, we recommend using within 1 week.

    Q. How long is my VivoTag labeled protein or antibody good for?
    A. When stored at 4 °C and protected from light, typically labeled protein is good for 4 weeks. If long term storage is needed, we recommend storing at -20 °C.


    Conclusion

    In this review, we have attempted to cover all the aspects of protein quality control, from the necessary initial sample assessment to sample optimization. For each step, a set of relevant techniques has been suggested (Figure 1A). The first-line methods are essential and should be used systematically for a full quality control assessment. Different complementary methods can be added depending on the protein sample peculiarities and quality control requirements. The suggested approaches for first line assessment include the “basic requirements for evaluating protein quality” that have been recently proposed [10], but go significantly beyond them. We also suggest a sequential experimental work-flow, to be followed as a check-list in order to optimize the time and effort spent on each sample (Figure 1B). This work-flow elaborates the protein quality control and storage optimization steps of the general protein production/purification pipeline [10]. Overall, this global synthetic step-by-step overview should hopefully lead to better protein samples and therefore to better chances of success in downstream applications. In line with community-based efforts that have been deployed in other fields like structural biology [69,70], proteomics and interactomics [71-74] or quantitative real-time PCR [75,76], research relying on purified proteins would gain significant reliability and credibility from the implementation of good practices, such as the systematic and transparent reporting of the results of purified protein quality control assessments, at least in the supplementary information sections of scientific publications.


    Watch the video: Πρωτεΐνες η αλήθεια (July 2022).


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