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9.6: Conjugation - Biology

9.6: Conjugation - Biology


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There are two other processes that can lead to horizontal gene transfer in bacteria: conjugation and transduction. In contrast to transformation, these processes “force” DNA into what may be a reluctant cell. In the process of conjugation, we can distinguish between two types of bacterial cells (of the same species). One contains a plasmid known as the sex factor (F) plasmid. These are known as a Hfr (high frequency recombination) cells. This plasmid contains the genes needed to transfer a copy of its DNA into a cell that lacks an F-plasmid, a so called F–cell. Occasionally, the F-plasmid can integrate (insert) itself into the host cell chromosome and when this happens, the F-plasmid mediated system can transfer host cell genes (in addition to plasmid genes) into an F–cell. To help make things a little simpler, we will refer to the Hfr cell as the DNA donor and F–cells as the DNA recipients.

To initiate conjugation, the Hfr cell makes a physical bridge to the F–cell. A break in the donor DNA initiates a process by which single stranded DNA is synthesized and moved into the recipient (F–) cell. The amount of DNA transported is determined largely by how long the transporter bridge remains intact. It takes about 100 minutes to transfer the entire donor chromosome from an Hfr to an F–cell. Once inside the F–cell, the DNA is integrated into the recipient’s chromosome, replacing the recipient’s versions of the genes transferred (through a process known as homologous recombination). Using Hfr strains with integrated F–plasmids carrying different alleles of various genes, and by controlling the duration of conjugation (separating the cells by placing them in a kitchen blender), experimenters were able to determined the order of genes along the chromosome. The result was the discovery that related organisms had the same genes arranged in the same order. The typical drawing of the circular bacterial chromosome is like a clock going from 0 to 100, with the genes placed in their respective positions, based on the time it takes to transfer them (in minutes). This is an example of synteny, that is the conservation of gene order along a chromosome266. We will return to synteny soon.

If the entire F-plasmid sequence is transferred, the original F–cell becomes an Hfr cell. If the Hfr cell loses the F-plasmid sequence, it reverts to a F–state. The end result of the conjugation process is similar to that obtained in sexual reproduction in eukaryotes (see below), namely the original F–cell now has a genome derived in part from itself and from the “donor” Hfr strain cell.


Using the Ka values in Table 9.6, calculate the pH of a buffer that contains the given concentrations of a weak acid and its conjugate base. a.0.55 M CH3COOH and 0.55 M NaCH3COO

Using the Ka values in Table 9.6, calculate the pH of a buffer that contains the given concentrations of a weak acid and its conjugate base.

a.0.55 M CH3COOH and 0.55 M NaCH3COO

Buffer Weak Acid Conjugate Base Ka
Acetic acid/acetate CH3COOH CH3COO − 1.8 × 10 − 5
Bicarbonate/carbonate HCO3 CO3 2 − 5.6 × 10 − 11
Dihydrogen phosphate/ hydrogen phosphate H2PO4 HPO4 2 − 6.2 × 10 − 8
Hydrogen phosphate/ phosphate HPO4 2 − PO4 3 − 2.2 × 10 − 13


Materials|methods

Cells of B. truncatella were obtained from Carolina Biological Supply Company (Burlington, NC, USA) and cells of C. magna were obtained from the American Type Culture Collection (#50128). Clonal cultures were established and grown in Volvic water with wheat grains and Klebsiella sp. B. truncatella cultures also included Paramecium sp. Individual cells were picked with a pipette and washed three times in sterilized Volvic water, then allowed to starve for 48 h. Ten starved cells from each species were individually whole genome amplified with REPLI-g Mini Kit (Hilden, Germany) following manufacturer’s instructions. For each species, the ten whole-genome amplified products were combined in equal DNA concentrations.

Amplified DNA from B. truncatella was sequenced with Illumina MiSeq v2 chemistry (13,288,644 2 × 250 bp reads) and Illumina HiSeq v3 chemistry 113,546,269 2 × 150 bp reads). Amplified DNA from C. magna was sequenced with MiSeq v3 chemistry (18,770,554 2 × 300 bp reads) and HiSeq v3 chemistry (28,298,554 2 × 150 bp reads). As the genome sizes of these two species are unknown, we could not estimate sequencing coverage. The optimal k-mer length for genome assembly was searched within a 21–201 range with KmerGenie v1.6976 ( Chikhi and Medvedev 2014), using the “diploid” parameter. Genomes were then assembled with Minia v2.0.7 ( Chikhi and Rizk 2012), setting the kmer minimal abundance to 5. The obtained contigs were then analyzed with AUGUSTUS v2.7 ( Stanke et al. 2004) for a structural annotation, using the following parameters: search on both strands genome is partial predict genes independently on each strand, allow overlapping genes on opposite strands report transcripts with in-frame stop codons species set to the ciliate T. thermophila. Reads were deposited in GenBank’s Sequence Read Archive under BioProject numbers PRJNA381863 and PRJNA382551.

A query database of 11 meiosis-specific genes and 40 meiosis-related genes from ciliate and nonciliate eukaryotes was established using literature and keyword searches of the NCBI protein database was taken from Chi et al. (2014a). For REC8, we used canonical eukaryotic sequences and T. thermophila’s noncanonical REC8 ( Howard-Till et al. 2013).The ORFs of the two colpodeans were searched by the query database using BlastP ( Altschul et al. 1990) and HMMER v3.0 ( Eddy 2001). Hits with E-values <10 −4 for the full sequence were retained. Verification of candidate homologs used reciprocal BlastP search against the nonredundant protein sequence database of NCBI ( supplementary Files 1 and 2 , Supplementary Material online).

In order to determine the strength of purifying selection acting on the inventoried meiotic genes, we measured ω = dN/dS, the number of nonsynonymous substitutions per nonsynonymous site divided by the number of synonymous substitutions per synonymous site. Sequences generated from this study were aligned with homologous sequences identified from T. thermophila, P. tetraurelia, Ichthyophthirius multifiliis, and Oxytricha trifallax by Chi et al. (2014a). Sequences were aligned in Geneious v4.8.3 ( Kearse et al. 2012) using Translation Align with ClustalW v2 ( Larkin et al. 2007). Sequences that did not have sufficient overlap with the other genes, were unalignable, or were determined to be paralogous to the genes from these other species were excluded from further analysis ( supplementary table 1 , Supplementary Material online). Maximum Likelihood genealogies of each gene were inferred with MEGA7 ( Kumar et al. 2016), and synonymous and nonsynonymous substation rates were estimated with PAML v4.8 ( Yang 2007). ω was first calculated between B. truncatella and C. magna. Then all species were included to test whether the lineage leading to C. magna exhibited higher values of ω, which would be expected if these genes were no longer functional and thus experiencing relaxed selection. Using codeml, we compared a model of evolution with one value of ω for the whole tree (model = 0) to a model where the C. magna lineage had a separate value of ω (model = 2). A log-likelihood ratio test was used to determine whether the second model provided a significantly better fit to the data.


Coccolithophores such as Emiliania huxleyi shown here are single-celled marine phytoplankton that produce calcium carbonate scales (coccoliths). They occur in all of the world’s oceans and together represent the largest source of biogenic calcium carbonate on Earth, contributing significantly to the global carbon cycle. Coccoliths, which are produced inside the cells, are secreted to the cell surface. The discovery of proton channels in the coccolithophore cell membrane provides new insight into how they cope with excess protons generated during calcification and will inform future studies of how these important organisms will respond to ocean acidification (see Taylor et al., e1001085).

Image Credit: Alison R. Taylor (University of North Carolina Wilmington Microscopy Facility)

Citation: (2011) PLoS Biology Issue Image | Vol. 9(6) June 2011. PLoS Biol 9(6): ev09.i06. https://doi.org/10.1371/image.pbio.v09.i06

Published: June 28, 2011

Copyright: © 2011 Taylor. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.


Base ionization constant, Kb

Base ionization constant (Kb) – the equilibrium constant for the ionization of a base also called the base dissociation constant .

The general equation for the reaction of a base, B, with water:

B (aq) + H2O (l) ⇌ BH + (aq) + OH - (aq)

Bronsted-Lowry bases react with water to produce OH - (aq) ions and a conjugate acid, which together determine the acid–base properties of the aqueous solution.

Weak bases, like weak acids, form dynamic equilibria in aqueous solutions.

For the reaction of a generic base with water, the equilibrium law equation, K, is written as follows:

Since the concentration (density) of water is a constant, it can be incorporated into the value of K (just as it was in the equilibrium law equation for Ka).

This yields a new constant, Kb, called the base ionization constant:

Weak bases are the conjugate bases of weak acids.

For example, the ethanoate ion, C2H3O2 - (aq), is the conjugate base of ethanoic acid, HC2H3O2 (aq). Similarly, the hypochlorite ion, ClO - (aq), is the conjugate base for hypochlorous acid, HClO(aq).


Practical Tips of ELISA

Enzyme-linked Immunosorbent Assays (ELISAs) combine the specificity of antibodies with the sensitivity of simple enzyme assays, by using antibodies or antigens coupled to an easily-assayed enzyme. ELISAs can provide a useful measurement of antigen or antibody concentration. Being one of the most sensitive immunoassays, ELISA offers commercial value in laboratory research, diagnostic of disease biomarkers, and quality control in various industries.

There are different types of ELISA according to the analyte detection method. Each of them have its own protocol but the basic principle is similar. An ELISA is a generally a five-step procedure:
1) Antigen coating
2) Blocking all unbound sites to prevent non-specific absorption
3) Add analyte and incubation
4) Add labeled antibody and incubation
5) Add reagents for colouration/ luminescence, thus give a positive result.

We have already discussed the ELISA format and general protocol in ELISA Guide and ELISA Development Guide. Here we mainly discuss about some basic knowledge and practical tips during the operation process of ELISA. If you need more information about commercial ELISA kit, please visit our ELISA Kit Product.

2. ELISA Plate (Solid Phase)

There are varies types of solid phase that can be used for ELISA, such as membrane, well plate and beads. Their characterizations are described in the following Table 1.

Table 1. Types of Immunoassay Solid Phases

Materiala Binding Capacity Type of Interaction
Membrane
Nitrocellulose
PVDF
Nylon

High
High
High

Hydrophobic, Hydrophilic
Hydrophobic
Hydrophobic
Plates and tubes
Polystyrene
Polyvinyl

Low
Low

Hydrophobic
Hydrophobic
Beads
Polystyrene
Derivatized polystyrene
Microparticles

Moderate
High
High

Hydrophobic
Covalent, Hydrophobic, Hydrophilic
Covalent and Hydrophobic

Most commonly, ELISAs are performed in 96-well (or 384-well) plates. Most plates are either polystyrene or derivatives of polystyrene obtained by chemical modification or irradiation of the surface. The capture protein can be either passively absorbed on the surface of polystyrene plate or covalent coupled through modifications that leave amine or reactive groups such as maleimide, hydrazine, or N-oxysuccinimide groups on the surface (Figure 1). It is this binding and immobilization of reagents that makes ELISA so easy to design and perform. Having the reactants of the ELISA immobilized to the microplate surface makes it easy to separate bound from unbound material during the assay. This ability to wash away nonspecifically bound materials makes the ELISA a powerful tool for measuring specific analytes within a crude preparation.

Figure 1. The surface chemistry of polystyrene plate and immobilized protein.

Polystyrene will bind a wide variety of proteins in an increasing amount depending on their concentration in the coating solution. The specific and optimal amount needs to be determined for each protein. Carbohydrates and heavily glycosylated proteins do not adsorb well to polystyrene by the forces described above because they have very little ability to participate in hydrophobic interactions. In order to adhere these molecules, one must resort to the covalent linkages.

A number of modifications have been made to the polystyrene surface that allow for covalent linking of molecules to the plastic surface (Figure 1c). Maleimide groups react with a sulfhydryl forming a covalent link between the plastic surface and a protein or peptide. Hydrazine reacts with aldehydes generated by periodate oxidation of carbohydrates. N-Hydroxysuccinimide (NHS) reacts with amines on peptides or proteins. Peptides either through the COOH end by using a cross linker such as carbodiimide or through the amine by using a homobifunctional cross linker such as disuccinimidyl suberate (DSS). Table 2 shows recommended method to immobilize different antigens on polystyrene plate.

Table 2. Different antigens and their recommended immobilizing method.

Direct Adsorption Covalent Linkage
Proteins
Peptides longer than 15-20 amino acids
Small molecule epitopes attached to a protein
Bacteria and virus
Heavily glycosylated proteins
Proteins in the presence of detergents
Carbohydrates
Short Peptides
Lipids
DNA

3. Antigen Coating

A key feature of the plate based ELISA is that capture protein (antibody or antigen) can be attached to surfaces easily by passive adsorption or covalent linkage. This process is commonly called coating. Multiple factors affect the antigen coating process. The rate and extent of the coating mostly depends on the following factors:
? Diffusion coefficient of the attaching molecule
? Ratio of the surface area being coated to the volume of the coating solution
? Concentration of the antigen being adsorbed
? Temperature
? Time of adsorption

These factors are linked. It is most important to determine the optimal antigen coating concentration for each systems. Time and temperature are also important factors controlling the amount of protein adsorbed. A concentration range of 1–10 μg/mL of protein, in a volume of 50-100 μL, is a good guide to the level of protein needed to saturate available sites on a polystyrene plate. Commonly used coating solutions are: 50 mM carbonate, pH 9.6 20 mM Tris-HCl, pH 8.5 and 10 mM PBS, pH 7.2. The most thorough adsorption and lowest well-to-well variation occurs overnight (16–18 hours) at 4°C with the wells sealed to prevent evaporation. Adsorption time can be speeded up by incubation at room temperature for 4–8 hours or 37°C for 1-4h. There are many more variations, and ultimately, each scientist must titrate a particular antigen to obtain a standardized regime. After coating process, don’t forget to remove the coating solution and wash the plate by filling the wells with 200μl PBS for 3 times. Then remove the wash solutions by flicking the plate gently over a sink. Remove any remaining drops by patting the plate on a paper towel.

With any trouble of antigen coating, please visit our Microplate Coating Service, or ask for Free Consultant of Our Specialists.

4. Blocking Process

Coating of wells with the specific binding partner, either antigen or antibody, leaves unoccupied hydrophobic sites on the plastic. These sites must be blocked in order to prevent nonspecific binding of subsequent reactants. If this is not effectively accomplished, the assay will suffer from high background signal and lowered specificity and sensitivity (Figure 2). These blockers work by reducing non-specific binding to increase the signal-to-noise ratio. To prevent non-specific binding, blocking buffers are used after the solid-phase coating step to block any remaining open binding sites.

Blocking reagents are typically chosen in an empirical manner. The optimum blocker for one assay may not perform well in other assays. The two major classes of blocking agents that have been tested are proteins and detergents.

Figure 2. ELISA plate with and without blocking process.

Detergents come in three classes: nonionic, ionic, and zwitterionic. Both ionic and zwitterionic are not recommend to use as blocker because they disrupt the hydrophobic interactions that bind proteins coated to the surface of the plastic. Typically, detergents used as blocking reagents are non-ionic such as Tween 20 and Triton X-100. Detergents are considered temporary blockers they do not provide a permanent barrier to biomolecule attachment to the surface because their blocking ability can be removed by washing with water or aqueous buffer. Unlike non-ionic detergents, proteins are permanent blockers and only need to be added once after the surface is coated with the capture molecule. Some of the most commonly used protein blockers are listed in Table 3 as well as their advantages and disadvantages.

Table 3. Advantages and disadvantages of commonly used protein blockers.

Ideal blocking agents have the following characteristics:
? Effectively block nonspecific binding of assay reactants to the surface of the well
? Do not disrupt the binding of assay components that have been adsorbed to the well
? Act as a stabilizer (prevent denaturation) of assay reactants on the solid phase
? Do not cross-react with other assay reactants
? Do not possess enzymatic activity that might contribute to signal generation of the substrate or degradation of the reactants
? Perform consistently across various lots

5. Antibody Preparation

As the antibodies are cornerstone of ELISA test, the choice of antibodies is obviously of prime importance. The most frequently faced problem is how to choose an antibody, monoclone (MAb) or polyclone (PAb)? In general, a MAb is often chosen as the primary antibody to establish the highest level of specificity in an assay, and a PAb is chosen as the secondary antibody, to amplify the signal via multiple binding events. However, any combination can be used. All candidate antibodies must be tested together with the intended sample type in order to select the best performers.

Primary/secondary antibody should be diluted in blocking solution to help prevent non-specific binding. A concentration of 0.1-1.0 μg/ml is usually sufficient.

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Table 4. Key performance differences between MAb and PAb

Monoclonal antibodies (MAb) Polyclonal antibodies (PAb)
a. Generally produced in mice or recombinantly, these antibodies recognize a single epitope.
b. Since only one antibody molecule can bind to the antigen, the interaction is highly specific but can lack sensitivity.
a. Produced in goats, sheep, chicken, rabbits and other animals.
b. Polyclonal sera is a heterogepneous composite of antibodies with unique specificities and the concentration of specific antibody (PAb) is typically 50-200mg/mL.
c. PAbs are able to recognize multiple epitopes on any one antigen which makes them less sensitive to antigen mutational changes.
d. PAbs are useful when the nature of the antigen is not well known. However, their quantity is limited by the lifespan of the animal.

6. Sample Preparation

The samples contains the main analytes which we need to detect by ELISA. An excellent preparation of sample solution may improve the ELISA test quality. The sample type could be variety such as serum, plasma, urine, cell or tissue lysates, saliva, milk, cell culture supernatant etc. Each sample may need a specific preparation process. A general protocol for sample preparation as following:
a. Protein extract concentration is at least 1-2 mg/mL.
b. Cell and tissue extracts are diluted by 50% with binding buffer.
c. Samples are centrifuged at 10,000 rpm for 5 min at 4°C to remove any precipitate before use.

And for each sample details, see Table 5.

Table 5 Common used ELISA sample types and their treatments.

Samples Treatments
Serum Collect whole blood into a tube without additives Keep at room temperature for 20 minutes. Centrifuge 10 minutes at 3,000 rpm. Aliquot into small tubes and store at -80°C until use. Minimize freeze/thaw cycles.
Plasma Collect whole blood into an EDTA, Citrate or Sodium heparin tube Centrifuge 10 minutes at 3,000 rpm at 4°C Aliquot into small tubes and store at -80°C until use. Minimize freeze/thaw cycles.
Urine Collect urine without adding stabilizers. Centrifuge the samples hard (eg. 10,000 x g for 1 min or 5,000 x g for 2 min). Aliquot, quick freeze in dry ice/methanol bath, and store at -80°C until use.
Saliva Collect samples and centrifuge at 10,000 x g for 2 min at 4°C. Aliquot supernatant and store samples at -80°C. Minimize freeze/thaw cycles.
Cell lysates Place tissue culture plates on ice. Remove the media and gently wash cells once with ice-cold PBS. Remove the PBS and add 0.5 ml extraction buffer per 100 mm plate. Tilt the plate and scrape the cells into a pre-chilled tube. Vortex briefly and incubate on ice for 15-30 min. Centrifuge at 13,000 rpm for 10 min at 4°C (this creates a pellet from the insoluble content). Aliquot the supernatant into clean, chilled tubes (on ice) and store samples at -80°C, avoiding freeze/thaw cycles.
Tissue lysates Dissect the tissue of interest with clean tools, on ice preferably and as quickly as possible to prevent degradation by proteases. Place the tissue in round bottom microfuge tubes and immerse in liquid nitrogen to "snap freeze". Store samples at -80°C for later use or keep on ice for immediate homogenization. For a

7. Buffer Solution

There are 5 mainly buffer solutions used in ELISA test: coating buffer, blocking buffer, washing buffer, substrate buffer and stop buffer.

Coating buffer usually 0.05 M carbonate buffer with pH=9.6.

Coating buffer Blocking buffer Washing buffer Substrate buffer Stop buffer
0.05M carbonate buffer, pH=9.6 See Table3 0.01M PBS-Tween 20, pH=7.4 Phosphoric-citric acid buffer, pH=5.0 2M H2SO4
Na2CO3 1.59g NaCl 8g 0.2M Na2HPO4 25.7mL
NaHCO3 2.93g KH2PO4 0.5g 0.1M Citric acid 24.3mL
ddH2O 1000mL Na2HPO4 2.9g ddH2O 50mL
adjust pH to 9.6 Tween 20 0.5mL OPD(Substrate) 4mg
storage in 4°C ddH2O add to 1000mL 30% H2O2 0.015mL

8. Washing Steps

The incubations that are performed in an ELISA allow high-affinity specific interactions to form among reactants. By washing several times between each incubation, the excess reactants are diluted to an undetectable background level. In order to effectively dilute the excess reactants, it is necessary to wash 3–5 times after each incubation. It is also a good idea to allow a 5 to 10 minute soak with wash buffer at each wash step. If the wash steps are being performed by hand, tap out the excess wash buffer at each step by banging the plate upside down on dry paper towels. Do not allow the plate to dry for extended periods between wash steps as this can lead to a reduction of activity.

9. Data Analysis

The ELISA assay yields three different types of data output:

? Quantitative: ELISA data can be interpreted in comparison to a standard curve (a serial dilution of a known, purified antigen) in order to precisely calculate the concentrations of antigen in various samples.
? Qualitative: ELISAs can also be used to achieve a yes or no answer indicating whether a particular antigen is present in a sample, as compared to a blank well containing no antigen or an unrelated control antigen.
? Semi-Quantitative: ELISAs can be used to compare the relative levels of antigen in assay samples, since the intensity of signal will vary directly with antigen concentration.

ELISA data is typically graphed with optical density vs log concentration to produce a sigmoidal curve. Known concentrations of antigen are used to produce a standard curve and then this data is used to measure the concentration of unknown samples by comparison to the linear portion of the standard curve (Figure 3).


Methods

Antigens

Albumin from bovine serum (BSA) was natural purified protein obtained from Fluka, USA. Crude venom of cobra (Naja kaouthia) and green pit viper (Trimeresurus albolabris) were purchased from the Thai Red Cross Society, Thailand. Aflatoxin B1-BSA conjugated and soluble Aflatoxin B1 (Sigma, Germany) were prepared from Aspergillus flavus. Amylase enzyme Type XII-A (Sigma, Germany) was prepared from Bacillus licheniformis. Purified Chick Embryo Cell (PCEC) rabies vaccine strain flury LEP (Chiron Behring, India) were inactivated rabies viruses that were cultured in primary chicken fibroblast cell. Cholangiocarcinoma, cell line (KKU-100), which is an egg-proven Opisthorchis-associated cholangiocarcinoma derived from porta hepatic cells [68], was a gift from Dr. Banchob Sripa.

Construction of pMod1 phagemid vector

Phage 3.2 vector (Maxim Biotech Inc, USA) was used as a basis for the construction of pMod1 phagemid vector. The new vector contains SfiI and NotI restriction sites and includes a hexahistidine tag and Myc tag. Two oligonucleotides, Mod1up (5' TCG ACC CAT GGC TCG AGG CGG CCG CAC ATC ATC ATC ACC ATC ACG GGG CCG CAG GGC C 3') and Mod1dn (5' CT GCG GCC CCG TGA TGG TGA TGA TGA TGT GCG GCC GCC TCG AGC CAT GGG 3') were annealed and ligated into the phage 3.2 vector (Figure 1) for the generation of pMod1. The insertion was made at ApaI/SalI sites using T4 DNA ligase (NEB, USA). The finished pMod1 vector was amplified by transforming E. coli DH5αF', and the phagemid DNA was prepared by miniprep extraction kit (Qiagen, Germany). The integrity of the plasmid was confirmed by automated DNA sequence analysis (Macrogen, Korea).

Construction of human scFv phage library

Generation of scFv gene repertoire

The peripheral blood donations from one hundred and forty healthy, non-immunized donors were collected into four pools, according to different blood groups. These blood samples were tested negative and discharged from the Thai Red Cross Society blood donation unit in Nakhon Ratchasima province. Total RNA was prepared from the B lymphocytes and pooled together. Approximately 2 ml of blood samples from each donor were collected into four pools, according to different blood groups (blood group A, B, O and AB). Thus, a total of approximately 280 ml of blood sample was used. B-lymphocytes were isolated from peripheral blood by using Ficoll plaque reagent (Amersham, USA). Briefly, the diluted blood sample (1:1 of blood per PBS) was carefully layered on top of the Ficoll plaque reagent, and then the two-phase solution was centrifuged at 400 × g for 30 minutes. B-lymphocytes were collected from the interface between the two phases. The interface contamination such as platelets and plasma proteins were removed by washing with PBS. Total RNA was extracted from B-lymphocytes by TRIzol reagent (Invitrogen, USA). B-lymphocytes were resuspended in 1 ml TRIzol and incubated at 65°C for 15 minutes with occasional inversion of the tube. After adding 0.2 ml of chloroform, the tube was vortexed for 15 seconds and then centrifuged at 12,000 × g for 15 minutes at 4°C. The aqueous phase was transferred to a new tube containing 1 μl of RnaseOut (40 U/μl, Invitrogen, USA), and 0.5 ml of isopropanol was added to precipitate RNA. The tube was incubated at room temperature for 10 minutes. The precipitated RNA was pelleted by centrifugation at 12,000 × g for 15 minutes at 4°C. The pellet was washed with 0.5 ml of 75% ethanol and then centrifuged at 12,000 × g for 15 minutes at 4°C. After the supernatant was removed, the pellet was air dried for 5 minutes at room temperature and dissolved in sterile deionized water. Then 1 μl of RnaseOut (40 U/μl, Invitrogen, USA) was added into total RNA and stored at -70°C. First strand cDNA was generated from 10 μg of total RNA, using MMuLV reverse transcriptase (NEB, USA) with a mix of 20 μM of oligo-dT18 and 8 ng of random hexamer primer. The genes for variable regions of heavy chain, κ light chain, and λ light chain (VH, Vκ, and Vλ) were amplified separately and recombined by three subsequent PCR reactions. The first set of PCR consists of 75 independent reactions to generate variable domains of the heavy and light chains. The heavy chain 5' primers were designed to include a SfiI site, and the light chain 3' primers include a NotI site. Light chain 5' primers were designed to include part of the linker region (Gly4Ser)3 and compatible with the heavy chain 3' primers (Table 1). Each variable region gene was separately amplified using hot start PCR in a reaction of 50 μl containing 5 μl cDNA and 1 μM of each 5' and 3' primer. This reaction was performed using the Taq polymerase (NEB, USA). The samples were heat at 94°C for 5 min, followed by 35 cycles of 94°C for 1 min, 55–65°C for 1 min, 72°C for 2 min. The final extension was performed at 72°C for 10 min. Equal amount of PCR products were pooled into collections of VH, Vκ, and Vλ gene repertoire, and purified from the low melting temperature agarose gel according to standard protocol [69]. In the second PCR, heavy and light chains were assembled and amplified using pfu DNA Polymerase (Promaga, USA). The assembly PCR reaction contained equal molar mixture of the pooled heavy (VH) DNA and pooled light (Vκ, or Vλ) gene repertoire. The assembly reaction was cycled 5 times (94°C for 45 s, 60°C for 50 s, and 72°C for 60 s) without primers. The third reaction created a full-length scFv gene repertoire from the second PCR by PCR amplification in the presence of pull-through primers. This PCR extended the scFv gene from the SfiI and NotI sites flanking scFv genes, using the following primers: PTfw (5'-CCT TTC TAT GCG GCC CAG CCG GCC ATG GCC-3') and PTrv (5'-CAG TCA TTC TCG ACT TGC GGC CGC ACG-3'). The reaction was performed using the Taq polymerase and 1 μl of assembled products from the second PCR. This pull-through PCR was cycled 30 times (94°C for 1 min, 60°C for 1 min, 72°C for 2 min), and a final extension at 72°C for 10 min. Then the samples were purified by a QIAquick PCR Purification Kit (QIAGEN, Germany) for the next step.

Cloning of the scFv into pMod1 vector

The amplified scFv DNA and pMod1 phagemid vector were sequentially digested with NotI and SfiI, by incubating at appropriate conditions for 10 hrs. The cut vector was then de-phosphorylated with calf intestinal phosphatase (NEB, USA) and gel-purified using Wizard ® DNA Clean-Up System (Promega, USA) before ligation. A total of 2.8 μg of digested scFv DNA were ligated into 5.5 μg of pMod1 phagemid vector, at a vector: insert molar ratio of 1:3, to generate the scFv-gene III fusion library. The ligation was done using T4 DNA ligase (NEB, USA) overnight at 16°C. Ligated DNA populations were electroporated into Escherichia coli (E. coli) TG1 (Maxim Biotech Inc, USA) using an Eppendrof 2510 electroporator (Eppendrof, USA). The transformed cells were then incubated for 1 hour at 37°C before spreading on TYE plate containing ampicillin (100 μg/mL) and glucose (1% w/v), the plate was incubated overnight at 37°C. Complexity of the library was determined at this step by serially diluting the transformed cells and counting the number of colonies. Ligation efficiency was also determined by counting the number of colonies from no-insert ligation. Colonies were then collected, mixed with glycerol, and stored at -80°C. The library stock was grown to log phase and rescued with M13KO7 helper phage (Maxim Biotech Inc, USA). Recombinant phage preparations were purified and concentrated by polyethylene glycol (PEG) precipitation before keeping at -80°C.

Determination of library size

A total of 8.3 μg of DNA was electroporated in to E.coli TG1 to generate the scFv phage library. After electroporation the cuvette was flushed with 6 ml of SOC medium and transformed cells were incubated for 1 hour at 37°C before spreading on TYE plate containing ampicillin (100 μg/mL) and glucose (1% w/v), then the plate was incubated overnight at 37°C. Complexity of the library was determined by serially diluting the transformed cells and counting the number of colonies. A volume of 100 μl from transformation reactions was taken and a four step 10-fold serial dilution was made. The 100 μl of each dilution was plated out. The appearance of 250 individual colonies from 10 4 dilution titer indicated a library diversity of 1.5 × 10 8 .

Selection of phage antibody library

Selection of phage particles displaying specific scFv fragments were performed on Immuno 96 MicroWell™ Plates (Nunc, Denmark). The different protein antigens (50–600 μg/ml) in phosphate-buffered saline (PBS) (or 0.1 mM NaHCO3, in case of amylase) were coated on the plates overnight at 4°C (in case of Rabies, the preparation was first incubated at 37°C for 2 hrs). For cholangiocarcinoma cell surface, approximately 5 × 10 5 cells growing in a 5-ml flask was used. Following blocking with 2% (w/v) skimmed milk powder in PBS (2% MPBS), a library containing between 10 11 and 10 12 phage particles were added and the plate was incubated for 2 hours at room temperature (RT 25–28°C). Non-bound phages were eliminated by washing 10–20 times with PBS containing 0.1% Tween 20 (PBS-T), followed by 10–20 times washing with PBS. The bound phages were eluted by incubation with 50 μl of 1 μg/μl trypsin for 10 min, followed by 50 μl of 50 mM glycine- HCl pH 2.0 (immediately neutralized with 50 μl of 200 mM NaHPO4, pH7.5 after 10 min). Eluted phages were used to infect exponentially growing E.coli TG1 cells by incubating for 30 min at 37°C. Infected cells were spread on TYE plate containing ampicillin (100 μg/mL) and glucose (1% w/v), then the plate was incubated overnight at 37°C. Individual phage-infected colonies were picked and grown for production of phagemid particles in 96-well plate. The culture was rescued using either M13KO7 or KM13 helper phage (MRC HGMP Resource Centre, Cambridge, UK) as describe elsewhere [55]. Rescued phage particles were used to test their antigen recognition properties by ELISA or to initiate subsequent rounds of selection using the similar conditions. Between one and two rounds of selection were performed for each antigen.

ELISA screening of selected clones

In order to detect antigen recognition, Immuno 96 MicroWell™ Plates were coated with approximately 5–200 μg/ml of each antigen. For cholangiocarcinoma, the cells were fixed with 4% paraformaldehyde in 96-well tissue culture plate. After overnight incubation at 4°C, plates were blocked with 2% MPBS for 1 hour at RT followed by three washes with PBS. The selected phage preparation was diluted 1:2 in 4% MPBS before adding into each well, and incubated for 1 hour at RT. The plates were washed three times with PBS-T, followed by three times with PBS, and incubated with a 1:5,000 dilution of a mouse anti-M13 phage-horseradish peroxidase (HRP) conjugate (Amersham-Pharmacia Biotech, Sweden) in 2% MPBS. The plates were washed again as described earlier. The ABTS (2,2-azino-di-3-ethyl-benzthiazoine-6-sulfonate) peroxidase substrate (Fluka, USA) was added, and the absorbance was read at 405 nm, using a Sunrise absorbance reader (TECAN, Austria).

Inhibition ELISA

The inhibition ELISA was performed as described in the normal ELISA method, except that the phage particles were pre-incubated in the presence of increasing amount of soluble Aflatoxin B1 from 0.039–5.0 μg/ml.

DNA fingerprint analysis and DNA sequencing

The diversity of the selected scFv clones was analyzed by comparing restriction enzyme digestion patterns, a procedure called DNA fingerprinting. The scFv sequence of individual clones was amplified by PCR using the following primers: PTfw (5'-CCT TTC TAT GCG GCC CAG CCG GCC ATG GCC-3') and PTrv (5'-CAG TCA TTC TCG ACT TGC GGC CGC ACG-3'). The amplified product was digested with a frequent cutting enzyme, BstNI (NEB, USA) and analyzed on 2% agarose gels. For DNA sequencing, plasmid DNA from different clones was purified using MiniPreps kit (QIAGEN, Germany) and the inserts were sequenced using the dideoxynucleotide chain-termination method with -96gIII primer (5'-CCC TCA TAG TTA GCG TAA CG-3'), corresponding to the vector sequence downstream of the scFv gene. After the amino acid sequences were translated, they were analyzed by using IgBLAST [36] and V BASE [34] software.

Soluble scFv antibodies production

E. coli HB2151 (Maxim Biotech Inc, USA) cells carrying the phagemid encoding the scFv antibody were grown in 10 ml of 2 × TY medium containing ampicillin (100 μg/mL) and glucose (1% w/v). After reaching an OD600 of 0.9, cells were harvested and grown at 30°C in glucose free 2 × TY medium containing 100 μg/mL ampicillin and 0.1 mM isopropyl-μ-d-thiogalactopyranoside (IPTG). The cell culture supernatant containing scFv were collected after induction for 20 hours. In some case, the cell lysate containing scFv were extracted after induction for 6 hours.


Absolute Quantification of Drug Vector Delivery to the Cytosol

Macromolecular drugs inefficiently cross membranes to reach their cytosolic targets. They require drug delivery vectors to facilitate their translocation across the plasma membrane or escape from endosomes. Optimization of these vectors has however been hindered by the difficulty to accurately measure cytosolic arrival. We have developed an exceptionally sensitive and robust assay for the relative or absolute quantification of this step. The assay is based on benzylguanine and biotin modifications on a drug delivery vector of interest, which allow, respectively, for selective covalent capture in the cytosol with a SNAP-tag fusion protein and for quantification at picomolar sensitivity. The assay was validated by determining the absolute numbers of cytosolic molecules for two drug delivery vectors: the B-subunit of Shiga toxin and the cell-penetrating peptide TAT. We expect this assay to favor delivery vector optimization and the understanding of the enigmatic translocation process.

Biological macromolecules such as peptides, proteins or oligonucleotides hold great therapeutic potential. They enable indeed to target “undruggable” proteins which lack cavities for small molecule binding, or to stimulate the immune system by antigen cross-presentation. However, macromolecules do not readily cross membranes. Most remain trapped at the plasma membrane or in intracellular compartments and do not reach their targets in the cytosol. Cytosolic arrival, via direct translocation across the plasma membrane or endocytosis followed by endosomal escape, is currently one of the main bottlenecks for the development of new macromolecular therapeutics. 1-3

For the optimization of cytosolic delivery, the process must be quantified. 4 Existing assays for this have a number of limitations, including the failure to distinguish between cytosolic versus intraluminal localizations and a lack of sensitivity and robustness (Table S1). Of note, two recently developed assays, the Chloroalkane Penetration Assay 5 and the NanoClick assay 6 rely on the HaloTag protein, 7 which covalently reacts to chloroalkanes. The cytosolic localization of the HaloTag reporter protein ensures the cytosolic specificity of the assay. Both assays measure the non-reacted reporter protein fraction, which is inversely proportional to the extent of translocated molecules. This limits the assay sensitivity. Furthermore, these assays do not provide absolute quantification of translocated molecules. Therefore, the quantification of small amounts of molecules reaching the cytosol remains challenging.

The example of siRNAs for the downregulation of disease-related proteins might be chosen to illustrate this point. For an efficient therapeutical effect, it is estimated that 2000–4000 siRNA molecules need to reach the cytosol per cell. 8, 9 The sensitivity of a cytosolic arrival assay is thus essential to detect such low numbers of molecules per cell and to optimize existing or develop new delivery tools.

To address this challenge, we have designed the Cyto-SNAP assay. This assay is based on a cytosolic capture protein assembled from SNAP-tag and mNeonGreen. The SNAP-tag reacts covalently with the small molecule benzylguanine (BG). 10 A vector of interest (e.g., protein- or peptide-based drug delivery tools) modified with BG will react with the SNAP-tag only if the vector reaches the cytosol (Figure 1 a).

A robust, sensitive, and quantitative cytosolic arrival assay. a) Schematic representation of the Cyto-SNAP assay. Upon membrane translocation, the BG-modified vector encounters the cytosolic mNeonGreen-SNAP-tag protein with which it reacts covalently. mNeonGreen is exploited for immunoprecipitation on beads coated with anti-mNeonGreen nanobodies, and the biotin moiety for ELISA. b) Parental, polyclonal mNeonGreen expressing (NG) or monoclonal mNeonGreen-SNAP-tag expressing (NG-SNAP) HeLa cell lines were treated with fluorescent SNAP-tag ligand BG-647-SiR. SiR fluorescence was only observed on NG-SNAP cells, in which it was homogeneously distributed in the cytosolic space. c) Demonstration of the high efficiency of the SNAP-tag reaction. Lysate from NG-SNAP cells was incubated with excess benzylguanine–biotin (BG–biotin) ligand, followed by streptavidin pull-down. Western blotting analysis showed that the cell lysate was depleted of mNeonGreen-SNAP-tag protein, which was indeed recovered on beads. d) Demonstration that the mNeonGreen immunoprecipitation (IP) is complete. The mNeonGreen-SNAP-tag protein was totally recovered on mNeonGreen-Trap beads, which confirmed the efficacy of the immunoprecipitation. e) Sensitivity and linearity of the assay. Known amounts of STxB-BG-biotin conjugate C were added into NG-SNAP cell lysate, followed by incubation at 37 °C, mNeonGreen-SNAP-tag immunoprecipitation and ELISA development. The obtained standard curve was linear over a wide range of concentrations. Even low picomolar STxB-BG-biotin concentrations were robustly detected. f) Demonstration that non-reacted SNAP-tag protein is efficiently quenched before cell lysis. Intact NG-SNAP and NG cells were incubated for 30 min at 4 °C or 37 °C in presence or absence (Ø) of SNAP-Cell® Block reagent. The cells were then washed, lysed, and lysates were incubated at 4 °C with or without STxB-BG-biotin conjugate A. Both 37 °C and 4 °C block conditions gave ELISA signals comparable to background signal without STxB-BG-biotin incubation.

mNeonGreen, 11 the second part of the cytosolic capture protein, is used for high affinity immunoprecipitation on beads coated with anti-mNeonGreen nanobodies (low Kd of 2 n m ). In this way, BG-tagged vector that reacts with the SNAP-tag of the cytosolic capture protein can be isolated. Finally, a biotin conjugated to the vector serves for quantification by ELISA (Figure S1).

We chose to validate the assay using two different vectors: i) Residues 47–57 from the HIV TAT protein, 12 which is one of the best-studied cell-penetrating peptides, and ii) the B-subunit of Shiga toxin (STxB), a vector for cancer cell targeting and immunotherapy. 13 After binding to its receptor, the glycosphingolipid Gb3, STxB is internalized and follows the retrograde transport route to the endoplasmic reticulum. STxB was shown to be able to translocate to the cytosol 14 and to efficiently deliver antigens for antigen cross-presentation, 13 making it an interesting model for a cytosolic arrival assay.

A monoclonal HeLa cell line, termed NG-SNAP, stably expressing the cytosolic mNeonGreen-SNAP-tag capture protein was generated for the assay. Diffuse mNeonGreen fluorescence and BG-fluorophore labeling were observed, which documented the expression of properly folded protein in the cytosol (Figure 1 b). To test whether the cytosolically localized mNeonGreen-SNAP-tag was fully functional, immunoprecipitation and pull-down experiments were performed against each of its two subunits: i) Lysates from NG-SNAP cells were incubated with an excess of BG–biotin, followed by streptavidin pull-down. This led to the complete depletion of mNeonGreen-SNAP-tag from the lysates, thereby proving that the SNAP-tag subunit was fully functional (Figure 1 c). ii) The same approach using anti-mNeonGreen nanobody beads also led to the depletion of mNeonGreen-SNAP-tag from the lysates, which further validated the functionality of the fusion protein (Figure 1 d). The combination of highly efficient SNAP-tag reaction and complete mNeonGreen immunoprecipitation laid the foundation for a fully quantitative assay.

Linearity and sensitivity are key parameters of a quantitative assay. To this end, low picomolar concentrations of BG- and biotin-tagged STxB were added to mNeonGreen-SNAP-tag-containing cell lysate, and incubated at 37 °C to allow for BG-SNAP-tag reaction. Subsequent mNeonGreen immunoprecipitation and ELISA development resulted in a linear standard curve, which documented the exquisite sensitivity of the assay (Figure 1 e).

Importantly, on cells, using a membrane-permeable BG-derivative, it was possible to quench non-reacted SNAP-tag prior to lysis (Figure 1 f). In the real assay format, this step is required to avoid post-lysis reaction between non-cytosolic BG-tagged vector and non-conjugated SNAP-tag in the lysate. Finally, STxB showed normal intracellular trafficking in NG-SNAP cells and in cells stably expressing mNeonGreen in the absence of SNAP-tag (termed NG cells), which were used as controls (Figure S2). All these findings qualified the engineered NG-SNAP and NG cell lines for the Cyto-SNAP assay.

The Cyto-SNAP assay (Figure S3) was first used to provide relative quantifications of cytosolic arrival between different conditions (incubation times, vector concentrations or inhibitor treatments). STxB (Figure 2 a,b) and TAT (Figure 2 g,h) translocation to the cytosol increased in a time- and concentration-dependent manner. For STxB, the translocation signal started to plateau at concentrations above 100 n m (Figure 2 b and S4a), while TAT translocation continued to increase exponentially beyond at least 20 μ m (Figure 2 h and S4b), suggesting that the process was receptor-independent for TAT, and receptor-dependent for STxB. Depletion of the STxB receptor, glycosphingolipid Gb3, by incubation of NG-SNAP cells with a glucosylceramide synthase inhibitor indeed reduced the cytosolic arrival of STxB to background level (Figure 2 c). Background was defined throughout all experiments as ELISA signal observed from NG cells. STxB translocation to the cytosol was also greatly impacted when incubations were performed at low temperatures, i.e., 4 °C or 19.5 °C (Figure 2 d). This observation was likely caused by changes in membrane organization leading to the disappearance of domain boundaries at which membranes are more permeable. 15, 16 In ATP depleted cells, STxB arrival in the cytosol was also strongly decreased (Figure 2 e). As for the 4 °C condition, endocytosis of STxB is inhibited in ATP-depleted cells. 17 Thus, it can be concluded that cellular entry is required for efficient translocation to the cytosol. The arrival of STxB in the cytosol was also found to be partially reduced when endosomal acidification was inhibited (Figure 2 f). Finally, TAT was compared to TAT-PEG6-GFWFG, previously reported to have an enhanced capacity to translocate to the cytosol. 18 We found no difference in cytosolic arrival at concentrations below 10 μ m (Figure 2 i). In contrast, TAT-PEG6-GFWFG translocated much more efficiently than TAT at concentrations above 15 μ m (Figure 2 i). TAT undergoes endocytosis at low concentrations, whereas at concentrations above 10 μ m direct translocation across the plasma membrane becomes the dominant mechanism (ref. 19 for a review, see ref. 20 ). It therefore appears likely that the strongly enhanced cytosolic arrival of TAT-PEG6-GFWFG at concentrations above 15 μ m resulted from direct translocation across the plasma membrane. These results qualified the Cyto-SNAP assay for membrane translocation measurements with different types of vectors.

Relative quantifications of STxB and TAT translocation to the cytosol, using the Cyto-SNAP assay on NG-SNAP cells. In all experiments, NG cells served as background controls. The cytosolic signal is expressed as percentage of the maximum signal obtained in each individual replicate experiment. a) Time dependency. Cells were incubated with 40 n m STxB-BG-biotin conjugate A for 0, 4, or 24 h (continuous incubation). The cytosolic signal increased with time. b) Concentration dependency. Cells were incubated for 4 h with STxB-BG-biotin conjugate A at the indicated concentrations. The cytosolic signal increased with concentration. c) Glycosphingolipid dependency. Cells were pre-treated or not for 5 days with the glycosylceramide synthase inhibitor Genz-123346, and then incubated for 4 h with 40 n m STxB-BG-biotin conjugate A. The translocation process to the cytosol was glycosphingolipid dependent. d) Temperature dependency. Cells were incubated for 4 h with 40 n m STxB-BG-biotin conjugate A at the indicated temperatures. Translocation to the cytosol was blocked at 4 °C and 19.5 °C. e) ATP dependency. Cells were incubated for 90 min with 40 n m STxB-BG-biotin conjugate A under ATP depletion conditions. Translocation to the cytosol was ATP dependent. f) Acidification dependency. Cells were incubated for 4 h with 40 n m STxB-BG-biotin conjugate A in the presence or absence of 100 n m of the V-ATPase inhibitor bafilomycin A1. Translocation to the cytosol was partly inhibited by bafilomycin A1 treatment. g) Time dependency of TAT translocation to the cytosol. Cells were incubated with 200 n m BG-biotin-TAT conjugate C for 0, 4, or 24 h (continuous incubation). The cytosolic signal increased with time. h) Concentration dependency of TAT translocation to the cytosol. Cells were incubated for 4 h with BG-biotin-TAT conjugate C at the indicated concentrations. The cytosolic signal increased with concentration. i) Comparison of TAT versus TAT-PEG6-GFWFG. NG-SNAP cells were incubated for 4 h at 37 °C with different concentrations of TAT or TAT-PEG6-GFWFG. Cytosolic signal is normalized to signal from 25 μ m TAT. TAT-PEG6-GFWFG has an increased cytosolic translocation capacity compared to TAT at concentrations above 15 μ m . Statistical analysis: two-tailed paired t-test * p<0.05 ** p<0.01 and *** p<0.001.

For absolute quantification of cytosolic arrival, the conjugation of BG and biotin reporter moieties was optimized in order to achieve 1:1 molar ratios of both BG and biotin per vector (or per monomer in the case of homopentameric STxB). We synthesized several scaffolds, comprising each BG, biotin, and maleimide for conjugation to the vectors (Scheme 1 and Figure S5–7). All STxB-based conjugates showed intracellular retrograde trafficking to the Golgi, as for non-modified STxB (Figure S8). With increasing PEG linker sizes, STxB conjugates C and D showed higher ELISA signal intensity when added directly in cell lysate (Figure 3 a). This was likely due to reduced steric hindrance for streptavidin binding to biotin on STxB pentamer whose BG had already reacted with the SNAP-tag. The same effect was not observed on the monomeric TAT (Figure 3 b). In contrast, cytosolic arrival was reduced for both STxB and TAT with the increased PEG linker size of scaffold D (Figure 3 c,d). An effect of long PEG linkers on cytosolic arrival was previously reported for the cell-penetrating peptide TAT-PEG-GWWG: PEG12 or PEG18 diminished the translocation efficiency when compared to PEG6. 18 In order to balance sensitivity and translocation efficiency, we chose to work with the maleimide-BG-biotin scaffold C. The resulting STxB conjugate C had a membrane translocation capacity similar to conjugate A, for which BG and biotin were separately coupled to STxB.

Evaluation of the different STxB and TAT conjugates for the Cyto-SNAP assay. a,b) Comparison of the ELISA signal obtained with the different STxB and TAT conjugates directly added into cell lysate. 40 p m of each STxB conjugate or 100 p m of each TAT conjugate were added to NG-SNAP cell lysate and incubated for 1 h at 37 °C for the SNAP reaction to occur before mNeonGreen immunoprecipitation and ELISA development on beads. c,d) Comparison of cytosolic arrival signal from the different STxB conjugates (40 n m ) or TAT conjugates (200 n m ) after 4 h incubation at 37 °C with NG or NG-SNAP HeLa cells.

Different strategies used for biotin and BG conjugation to vectors a) Biconjugation method: Use of commercial NHS ester—BG for conjugation to amines of the vector, and maleimide-biotin for conjugation to thiol. b–d) Monoconjugation method: Synthesized maleimide-benzylguanine-biotin molecules for conjugation to thiol of the vector. Different PEG linker sizes were tested in order to avoid steric hindrance between vector, the BG reaction with the SNAP-tag, and streptavidin binding to the biotin moiety.

The choice of lysis conditions was also critical for quantification to ensure that vectors are extracted from membranes with which they may remain associated even after translocation to the cytosolic compartment (see ref. 14 for STxB). We used high salt concentrations and sonication as recommended for TAT extraction, 21 and optimized the choice of detergent to achieve complete STxB extraction (Figure S9a). These conditions remained fully compatible with the requirement for SNAP-tag reaction and immunoprecipitation (Figure S9b,c).

For absolute quantification, a standard curve is essential. For this, we have devised a simple approach (Figure S10): Known amounts of BG and biotin-tagged vector were mixed into NG-SNAP cell lysate, leading to complete reaction with the SNAP-tag protein of the lysate. As shown in Figure 1 e, low picomolar sensitivity was reached with this approach. Using the standard curve, the amount of vector translocated to the cytosol and the total amount of vector associated with NG-SNAP cells were then determined (Figure S10). For the total cell-associated amount, the SNAP-tag quenching step was omitted, such that BG and biotin-tagged vector molecules fully reacted with unquenched SNAP-tag in the cell lysate. Of note, immunoprecipitation and subsequent ELISA steps were performed for all samples in parallel under identical conditions such that absorbance readings could be compared directly.


Bacterial Conjugation Steps

In order to transfer the F-plasmid, a donor cell and a recipient cell must first establish contact. At this point, when the cells establish contact, the F-plasmid in the donor cell is a double-stranded DNA molecule that forms a circular structure. The following steps allow the transfer of the F-plasmid from one bacterial cell to another:

Step 1

The F + (donor) cell produces the pilus, which is a structure that projects out of the cell and begins contact with an F – (recipient) cell.

Step 2

The pilus enables direct contact between the donor and the recipient cells.

Step 3

Because the F-plasmid consists of a double-stranded DNA molecule forming a circular structure, i.e., it is attached on both ends, an enzyme (relaxase, or relaxosome when it forms a complex with other proteins) nicks one of the two DNA strands of the F-plasmid and this strand (also called T-strand) is transferred to the recipient cell.

Step 4


Watch the video: bio367-Genetics-Conjugation (October 2022).