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How do we regulate the production of proteins when designing plasmids?

How do we regulate the production of proteins when designing plasmids?



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I think it should be no surprise that I, as many others, am interested in the new COVID-19 vaccines being developed. In my region of the world there are two mayor candidates. One is mRNA based and one is, as far as I understand, plasmid based.

I have a passable understanding of the mRNA based vaccine. I also know what a plasmid is, and how it is transferred between cells, but my understanding is otherwise limited.

As far as I understand we want the plasmid to be "unlooped" and grafted onto our target cells DNA. This, as far as I reckon, should enable the cell to produce whatever protein the plasmid coded for. In vaccines this lets us device anti-measures (vague because of my lack of knowledge) against the virus… But, how do we regulate the production of these anti-measures? Can we include a "regulator" for the production in the plasmid, so that the modified cells only produce the proteins when the virus is present? Or do we just let the cells continuously produce the anti-measure proteins, regardless of their necessity?

My main interest here lies in where the permanency of the vaccines induced changes lie. With the mRNA vaccine I understand that the permanency lies in the regulated immune system, with other cells being otherwise unchanged after the mRNA decomposes. Regarding the DNA/plasmid vaccine I'm still unwise.

Thank you for your time, and happy holidays.


The Institute for Creation Research

The March 1998 Impact article"Cloning - What is It and Where is It Taking Us?" discussed the procedure of cloning by somatic cell transfer. In that procedure, the nucleus from a cell derived from an embryo, a fetus, or tissue of an adult is inserted into an egg from which the nucleus has been removed. After the egg develops to the appropriate stage, the embryo is inserted into the uterus of a properly prepared female and allowed to develop to term. This produces an offspring essentially genetically identical to the animal that provided the nucleus that was inserted into the egg. This article will discuss the transfer of genes from one species to another in order to endow the recipient species with beneficial properties, or to enable the recipient to produce human proteins for injection into human patients who lack a vitally important protein.

Production of Therapeutic Proteins by Gene Transfer

There are many proteins essential to good health that some people cannot produce because of genetic defects. These proteins include various blood-clotting factors causing hemophilia, insulin (resulting in diabetes), growth hormone (resulting in lack of proper growth), and other proteins, the administration of which corrects pathological conditions or results in other therapeutic benefits. The early work in this field employed bacteria. Some bacteria in a bacterial culture may contain small circular DNA molecules called plasmids. These plasmids are not part of the chromosomal DNA that is possessed by all the bacteria of the culture. As these exceptional bacterial cells reproduce by binary fission or cell division, the plasmids are transmitted to the daughter cells. They can also be transmitted to other cells by conjugation. Scientists have learned how to utilize plasmids to transfer human genes to bacterial cells. If the gene inserted into the plasmid of bacteria is the human gene for insulin, for example, the bacteria into which this gene is inserted produces human insulin.

Insertion of a DNA section into a plasmid

Scientists have identified and isolated enzymes (called restriction enzymes, or restriction endonucleases) each of which cuts genes in very specific places. More than 100 of these enzymes have been isolated. After the position of the human gene that codes for the desired therapeutic protein has been located on the chromosome, the gene is cut out using the appropriate restriction enzymes and the gene is isolated. The same restriction enzymes are used to cut out a piece of the circular DNA plasmid. Thus, the two ends of the human gene will be those that will link up with the open ends of the plasmid. An enzyme called DNA ligase is used to couple each end of the gene to the open ends of the plasmid, restoring a circular DNA molecule with the human gene replacing the piece cut out of the plasmid. These plasmids, now including the human gene, are reinserted into bacteria. These bacteria are cultured, producing large quantities of identical bacteria carrying the human gene that is reproduced along with the bacterial DNA. Furthermore, these bacteria produce the human protein coded for by the spliced human gene. The protein is isolated from the bacterial culture, purified, and injected into those patients suffering from pathological conditions because their bodies cannot produce sufficient quantities of the protein.

Before genetic engineering, these proteins had to be painstakingly isolated from tissues or blood, but since they are produced in such minute quantities, the isolation of significant quantities required the processing of large quantities of material. As a result, they were very expensive. Relatively much larger quantities were obtained from genetically altered bacterial cultures, but the cost, although less, was still high. The bioreactors (that is, the machinery required to culture large quantities of the bacteria containing the human gene) are enormously expensive and must be operated by several scientists and technical assistants. Furthermore, the proper operation of the bioreactor is sensitive to small changes in temperature and composition of the culture broth.

Fortunately, an alternative method has been developed which promises to be much less expensive and much more efficient. This method utilizes an animal, such as a pig, as the bioreactor. It required years for scientists to design, develop, and build the very expensive, difficult to operate bioreactor devised by man. God had already devised a much more efficient, much cheaper bioreactor. Scientists finally realized that it would be possible by genetic manipulation to induce a female pig, cow, or other animal to produce the desired human proteins in its milk.

Genetic engineering of a milk protein

This procedure has now been successful in both pigs and cows. Among animals, the pig has a number of advantages. Its gestation period is only four months. At 12 months of age the pig is fertile and produces large litter sizes (usually 10 to 12 piglets). A lactating pig produces 300 liters (about 315 quarts) of milk in a year. The procedure is carried out as follows:

  1. The DNA fragment (gene) that codes for the needed human protein is isolated.
  2. The DNA fragment that promotes production of proteins in mammary glands is isolated and linked or combined to the human gene.
  3. Fertilized eggs from a donor pig are obtained.
  4. The human DNA is injected into an egg in the region of the male pronucleus (DNA from the sperm before it unites with DNA of the egg) using a very slim micro pipette. The human DNA is thus incorporated into the pig nuclear DNA.
  5. The egg is implanted in the uterus of another pig and develops into a newborn female pig.
  6. The desired protein is isolated from the milk of the female pig once grown.

This procedure was carried out successfully by American scientists and the results were published in 1994. 1 The human gene they used codes for Protein C that acts to control blood clotting. They obtained one gram of Protein C from each liter of milk produced by the pig. This is 200 times the concentration found in human blood plasma. Only about one third of the Protein C obtained was biologically active. The reason for this is that many modifications of a protein must be performed in a cell after the protein chain is formed. For example, sections are cut out of the protein complex sugar groups may be attached at certain places in the protein chain and cell wall anchor groups may be added. The scientists discovered that a key processing enzyme, called furin, was present in insufficient quantities, so they added to their gene complex the gene that codes for furin. This increased the yield of active Protein C. The human proteins produced in this way must be tested for safety and effectiveness. At this writing, an anti-clotting protein called anti-thrombin is now being tested in clinical trials.

Comparing this procedure to the use of bioreactor machines illustrates the fact that a generation of biochemical engineers failed to match the abilities of a tool for making proteins (the pig) that God had prepared. The mammary gland is optimized to maintain a high density of cells to deliver to them an ample supply of nutrients and to channel proteins that are produced in a form that can easily be isolated and purified. This procedure has proven to be successful and promises to provide a method for producing valuable therapeutic proteins at much lower cost.


Applications of synthetic peptides

The invention of peptide synthesis in the fifties and sixties spurred the development of different application areas in which synthetic peptides are now used, including the development of epitope-specific antibodies against pathogenic proteins, the study of protein functions and the identification and characterization of proteins. Furthermore, synthetic peptides are used to study enzyme-substrate interactions within important enzyme classes such as kinases and proteases, which play a crucial role in cell signaling.

In cell biology, receptor binding or the substrate recognition specificity of newly discovered enzymes can often be studied using sets of homologous synthetic peptides. Synthetic peptides can resemble naturally occurring peptides and act as drugs against cancer and other major diseases. Finally, synthetic peptides are used as standards and reagents in mass spectrometry (MS)-based applications. Synthetic peptides play a central role in MS-based discovery, characterization and quantitation of proteins, especially those that serve as early biomarkers for diseases.

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Results and discussion

Engineering of LPS biosynthesis in E. coli

We began our construction of LPS-free E. coli K-12 using the Kdo-depleted strain KPM22 L11 [12]. This strain contains deletions of kdsD and gutQ, which encode d -arabinose 5-phosphate isomerases essential for the biosynthesis of Kdo [13,14], and a C:G to T:A transition at position 52 of msbA, which acts as a suppressor of the normally lethal ∆Kdo phenotype [12]. To produce a strain that contains lipid IVA as the only LPS outer membrane component and that cannot revert to synthesize endotoxic derivatives, we sequentially generated mutant strains with unmarked deletions of the genes lpxL, lpxM, pagP, lpxP and eptA. These genes encode enzymes acting downstream of Kdo incorporation into lipid IVA, and are either part of the constitutive pathway of Kdo2-hexaacyl lipid A biosynthesis (the Kdo2-lipid IVA lauroyl-ACP acyltransferase LpxL and the Kdo2-lauroyl-lipid IVA myristoyl-ACP acyltransferase LpxM), modify lipid A with additional acyl chains (the phospholipid:lipid A palmitoyl transferase PagP and the Kdo2-lipid IVA palmitoleoyl-ACP acyltransferase LpxP), or append phosphoethanolamine (P-EtN) under certain conditions (the lipid A P-EtN transferase EptA) [15] (Figure 1). Analysis of the LPS isolated from this strain, designated KPM318, using electrospray-ionization Fourier-transformed ion cyclotron mass spectrometry (ESI FT-ICR) revealed a single primary peak at 1404.85 u, consistent with the structure of the unmodified tetraacyl-1,4′-bisphosphate LPS precursor lipid IVA (Figure 2A).

Biosynthetic reactions targeted during the construction of E. coli strains KPM318 and KPM404. The late acyltransferases LpxL and LpxM of the constitutive pathway transfer laurate and myristate, respectively, to Kdo2-lipid IVA to form the characteristic acyloxyacyl units of hexaacylated Kdo2-lipid A. In contrast, LpxP, PagP and EptA are regulated in response to certain stimuli such as incorporation of palmitoleate in place of laurate by LpxP at low growth temperatures or PagP-catalyzed palmitoylation of lipid A upon phospholipid translocation to the outer leaflet of the outer membrane, for example, in strains defective in LPS biosynthesis. The acyl-acyl carrier protein (ACP) serves as the preferred acyl donor for various lipid A-acyltransferases.

Charge deconvoluted ESI FT-ICR mass spectra in negative ion mode of lipid IVA isolated from BW30270-derived mutants. The lipid IVA (calculated mass 1404.854 u) was extracted from E. coli (K-12) strains KPM318 (A) and KPM335 (B). Mass numbers given refer to the monoisotopic masses of neutral molecules. Peaks representing molecules with variations in acyl chain length are labeled with asterisks (∆m = 14.02 u). The molecular ion at 1484.82 u in panel B indicates the presence of a minor fraction of 1-diphosphate lipid IVA. The structures of lipid IVA and 1-diphosphate lipid IVA are shown as insets in panels A and B, respectively.

The successful construction of KPM318 demonstrated the viability of E. coli containing only lipid IVA. However, like other E. coli K-12 strains with partially defective outer membranes such as the ∆Kdo prototype strain KPM22 [11], KPM318 displayed growth defects at temperatures above 40°C. To overcome this, we isolated a series of stable temperature-resistant derivatives of KPM318 capable of growing exponentially at 42°C. KPM335 was the most robust of the spontaneous mutants. Whole-genome sequencing of KPM335 identified a single de novo mutation in comparison to the temperature sensitive KPM318 parent strain, a G:C to T:A transversion at base number 181 of the frr gene (Additional file 1: Table S1). The frr gene encodes an essential ribosome recycling factor which has been described to play multiple cellular roles, such as in disassembly of the post-termination complex [16], prevention of translation errors [17], promoting translational coupling [18], and increasing cell viability through polyamine stimulation of stationary-phase E. coli cultures [19]. The derivation of KPM335 argues strongly for a direct correlation between the development of the frr181 allele and the ability of the strain to tolerate the ∆Kdo phenotype at 42°C. However, elucidation of the underlying mechanism must await further investigations.

ESI FT-ICR analysis of the LPS isolated from KPM335 revealed no significant changes in the LPS composition, with lipid IVA remaining the predominant LPS-related molecule as in the parent KPM318 strain (Figure 2B). However, in contrast to KPM318, spectra of KPM335 displayed a minor peak with a molecular mass of 1484.82 u, consistent with the structure of 1-diphosphate lipid IVA. Touzé and coworkers have shown previously that transfer of a second phosphate group to the 1 position of lipid A is catalyzed by LpxT, an inner membrane protein of the undecaprenyl-pyrophosphate phosphatase family, which is able to phosphorylate Kdo2-lipid IVA in vitro, but not lipid acceptors lacking Kdo [20]. Thus, the presence of a minor tris-phosphorylated lipid IVA fraction in KPM335 argues against an absolute requirement of LpxT for Kdo-glycosylated lipid A acceptors under in vivo conditions. It remains unclear, however, why phosphorylation of lipid IVA, albeit with a very low efficiency, can occur in KPM335, but apparently not in its KPM318 parent strain.

Apart from the frr181 mutation in KPM335, a total of 12 mutations were specific to both KPM318 and KPM335, including ∆kdsD, ∆gutQ, ∆lpxL, ∆lpxM, ∆pagP, ∆lpxP, ∆eptA and msbA52 as a prerequisite for synthesis of lipid IVA as the predominant LPS-related outer membrane component. For another four mutations, we believe that they arose spontaneously during the generation of mutant strains, namely a silent mutation located in yqiI, two missense mutations in gor and waaY, and a point mutation in the non-coding region upstream of the deleted eptA gene. The latter mutation is most likely a result of KPM274 construction as a donor strain of the ΔeptA::kan cassette. The mutation is located within the sequence of the homology region of primer ECOeptAH1 (Additional file 2: Table S2) and suggests an error in PCR amplification of the kanamycin resistance cassette targeting the eptA gene and ultimately integration into the genome of KPM318. Compared to the reference genome sequence of the common E. coli MG1655 progenitor, the strains BW30270, KPM318 and KPM335 share sequence variations at six locations. Of these, both a silent nucleotide substitution at position 114 and a single nucleotide insertion at base number 253 of ylbE, and deletion of nucleotide 151 within the glpR gene have been described recently as genetic variations in common E. coli MG1655 stock cultures [21]. Furthermore, while E. coli MG1655 has been shown to express a defective ribonuclease PH due to a frameshift by deletion of the first nucleotide of codon 223 of the pseudogene rph-1 [22], insertion of a single nucleotide as the first base of codon 224 of rph-1 predicts reconstitution of RNase PH function and elimination of the polar effect of the rph-1 mutation on the downstream pyrE gene in BW30270, KPM318 and KPM335.

Next, we decided to replicate the unique set of unmarked genomic deletions in the genetic background of the popular E. coli expression strain BL21 (DE3). As a first step towards the construction of an LPS-free E. coli BL21 (DE3) derivative, we replaced the wild-type msbA gene with the msbA148 suppressor allele of strain KPM22 L1 [12]. This allowed the resulting strain, MWB03, to tolerate null mutations in otherwise essential genes of the LPS biosynthesis pathway. We then sequentially deleted the same genes that had been deleted during the creation of KPM318. Whole-genome sequencing of the final strain, KPM404, confirmed the presence of the msbA148 suppressor mutation (Additional file 3: Table S3), and verified the absence of the kdsD, gutQ, lpxL, lpxM, pagP, lpxP and eptA genes. In comparison to the genome sequence of E. coli BL21 (DE3), we further identified a silent mutation in yceJ, three missense changes within the coding sequences of YadG, ECD_00513 and RstA, and a point mutation in the intergenic region between nudC and hemE. Finally, a total of five single nucleotide substitutions were counted in the region between nucleotides 46 and 222 of the basR gene downstream of ∆eptA. These substitutions perfectly matched the sites of basR sequence variations in E. coli B and K-12, indicating that about one third of the BasR-encoding gene of KPM404 was replaced by the corresponding basR sequence of E. coli K-12. Just as for the construction of KPM318, the E. coli K-12 strain KPM274 served as the donor for transfer of the ΔeptA::kan cassette via P1vir transduction to yield strain KPM403, which well explains the generation of a basR hybrid sequence by co-transduction of the ΔeptA::kan cassette and adjacent basR sequences. In fact, as described above, the use of KPM274 as a ΔeptA::kan donor strain also explains why KPM404 carries a point mutation at the same position upstream of the deleted eptA gene as in KPM318.

Mass spectrometric analyses of the LPS profiles from mutants established in the BL21 (DE3) genetic background underscored the need for radical modification of LPS biosynthesis to accomplish synthesis of only lipid IVA in KPM404 (Figures 3 and 4). While disruption of the gutQ, kdsD and lpxL genes in the intermediate mutant strain KPM396 resulted in synthesis of non-glycosylated lipid IVA precursors lacking the secondary lauroyl and myristoyl chains, mass spectrometry revealed a rather heterogeneous mixture of differently modified lipid IVA species. The spectra displayed four prominent peaks with molecular masses of lipid IVA substituted with one P-EtN group (1527.86 u), lipid IVA modified with two P-EtN moieties (1650.87 u), and palmitoylated lipid IVA molecules carrying either one (1766.09 u) or two (1889.10 u) P-EtN residues. Since lipid A palmitoylation seems to be an indication for an adaptive response to aberrant translocation of phospholipids to the outer leaflet of the outer membrane [23], we suspect that PagP-mediated transfer of palmitate to lipid IVA is triggered by perturbations of outer membrane lipid asymmetry in Kdo-depleted strains of the KPM series. As shown for the LPS sample of KPM400, complete loss of the palmitoylated lipid IVA fraction was achieved by deletion of the pagP gene, leaving lipid IVA molecules modified with either one or two P-EtN groups unaffected.

Charge deconvoluted ESI FT-ICR mass spectra in negative ion mode of LPS isolated from BL21 (DE3)-derived mutants. Mass numbers given refer to the monoisotopic masses of neutral molecules. Peaks representing molecules with variations in acyl chain length are labeled with asterisks (∆m = 14.02 u). The mass spectra depict the progress in elimination of lipid IVA heterogeneity by sequential deletion of genes encoding the addition of acyl chains and P-EtN to the lipid A precursor.

Structures and molecular masses of lipid IVA molecules identified by ESI FT-ICR mass spectrometry in BL21 (DE3)-derived KPM mutants. The ESI FT-ICR mass spectra are shown in Figure 3. Modifications of lipid IVA with palmitate (green) and P-EtN (magenta) are catalyzed by PagP and EptA, respectively.

Due to the block in Kdo biosynthesis and the lack of LpxL, Kdo2-lipid IVA and Kdo2-(lauroyl)-lipid IVA, the preferred substrates for LpxP and LpxM, respectively [24-26], cannot be synthesized in strains derived from KPM396. In addition, previous work has shown that expression of LpxP is induced under conditions of cold shock (12°C) to incorporate an unsaturated C16:1 acyl chain at the expense of a laurate (C12:0), perhaps reflecting the demand to adjust membrane fluidity in the cold [25,27]. It was therefore not surprising that deletion of the lpxP and lpxM genes did not exhibit an obvious effect on lipid IVA composition of KPM400 and KPM402, respectively. There is no data indicating that LpxP and LpxM are capable of using lipid IVA as an acceptor substrate. It seems to be quite possible, however, that both enzymes show low levels of activity under specific conditions. Contrary to the proposed physiological role of LpxP in adaptation of E. coli cells to low growth temperatures, limited induction of LpxP expression has been demonstrated as a potential compensatory mechanism even at 30°C in lpxL and lpxL lpxM mutants of E. coli W3110 [28]. Likewise, LpxM was able to transfer a myristoyl chain directly to Kdo2-lipid IVA in E. coli W3110 strains lacking lpxL and lpxL lpxP [28].

Like PagP and LpxP, EptA-dependent modification of lipid A with P-EtN is part of the complex regulatory network associated with the structural redesign of LPS upon exposure to changing environmental conditions or envelope stress factors. As shown for E. coli K-12, modification of lipid A with P-EtN occurs under certain conditions such as in response to external stimuli like ammonium metavanadate [29] or mild acid pH [30]. Although P-EtN appears to be transferred predominantly to the 1-phosphate group of lipid A [31], double P-EtN substitutions at the 1- und 4′-phosphate positions were evident in lipid A of E. coli K-12 lacking LpxT activity [32] and lipid IVA of a mutant strain defective in MsbA-dependent translocation of LPS across the inner membrane [12]. Based upon ESI FT-ICR analysis presented here, deletion of the eptA gene was clearly necessary as well as sufficient to prevent lipid IVA of KPM404 from being substituted with one or two P-EtN residues. Thus, our data not only corroborate previous findings on the ability of EptA to transfer P-EtN to both the 1- and the 4′-phosphate group of lipid A [32], but also provide experimental evidence for its ability to use lipid IVA as a substrate for single and double P-EtN modifications.

In contrast to the E. coli K-12 strain KPM318, integration of lipid IVA into the outer membrane of KPM404 did not result in a temperature-sensitive phenotype of the BL21 (DE3)-derived mutant (data not shown). Although closely related at the genomic level [33], combined analysis of the genomes, transcriptomes, proteomes and phenomes of E. coli K-12 and BL21 (DE3) revealed significant differences in their cellular physiology and metabolism [34], which may explain the differences in the ability of KPM318 and KPM404 to maintain outer membrane integrity in the presence of lipid IVA at temperatures above 40°C.

Biological activity of engineered E. coli cells and LPS

To test the endotoxic potential of the engineered E. coli K-12 and B strains, we performed TLR4/MD-2 activation assays using HEK-Blue hTLR4 cells. Stimulation of these cells, which express human TLR4, MD-2 and CD14 on their surfaces, induces the production of the NF-κB- and activator protein-1 (AP-1)-dependent reporter secreted embryonic alkaline phosphatase (SEAP). The phosphatase levels can be determined by reading the absorbance at 655 nm using a colorimetric substrate. In order to address the question of whether NF-κB is specifically induced via the hTLR4/MD-2 signaling pathway, HEK-Blue Null2 cells, the parental cell line of HEK-Blue hTLR4 cells lacking the hTLR4/MD-2 receptor complex, were used as a control in all hTLR4/MD-2 activation assays. The strains KPM318, KPM335 and KPM404 as analyzed by challenging HEK-Blue hTLR4 cells with an increasing number of colony forming units (cfu) of up to 10 6 cfu/ml were virtually free of hTLR4/MD-2-stimulating activity, whereas their parental strains BW30270 and BL21 (DE3) elicited a substantial hTLR4/MD-2 activation already at 10 3 cfu/ml (Figures 5A, B, 6A and B). When the extracted LPS of the strains was subjected to the TLR4-specific assay, we could confirm the lack of endotoxic activity of the samples isolated from KPM318, KPM335 and KPM404 (Figures 5C, D, 6C and D). The data also demonstrated that palmitoylation of lipid IVA (when PagP was expressed) and/or modification of lipid IVA with one or two P-EtN groups (when EptA was expressed) in KPM396, KPM400 and KPM402 are capable of conferring certain hTLR4/MD-2 stimulatory activity on the otherwise endotoxically inactive tetraacylated lipid A precursor (Figure 6). Our results allow us to draw the major conclusion that inactivation of regulated lipid IVA modifications as demonstrated herein to be present in BL21 (DE3)-based intermediate mutant strains is a crucial prerequisite to yield consistently endotoxin-free E. coli strains.

Dose–response curves of NF-κB induction by whole bacterial cells and LPS of E. coli K-12 strains. The samples were assayed with HEK-Blue hTLR4 cells for hTLR4/MD-2-mediated NF-κB induction by colorimetric determination of NF-κB-dependent SEAP activity (A and C). HEK-Blue Null2 cells, the parental cell line of HEK-Blue hTLR4 cells lacking the hTLR4/MD-2 receptor complex, were used as a control (B and D). HEK-Blue hTLR4 and Null2 cells were stimulated with tenfold serial dilutions of whole bacterial cells (A and B) and LPS extracts (C and D) of KPM318 and KPM335 in comparison to their parental strain BW30270, respectively. The values represent the means and standard deviations from three individual experiments. In all experiments, assayed samples showed low level stimulation of HEK-Blue Null2 cells, indicating that NF-κB-dependent SEAP expression was specifically induced via the hTLR4/MD-2 signaling pathway in HEK-Blue hTLR4 cells.

Dose–response curves of NF-κB induction by whole bacterial cells and LPS of E. coli BL21 (DE3) strains. The samples were assayed with HEK-Blue hTLR4 (A and C) and Null2 (B and D) cells for relative NF-κB induction by colorimetric determination of NF-κB-dependent SEAP activity. Relative NF-κB induction was measured following stimulation of HEK-Blue hTLR4 and Null2 cells with tenfold serial dilutions of whole bacterial cells (A and B) and LPS extracts (C and D) of E. coli BL21 (DE3) and BL21 (DE3)-derived KPM mutants, respectively. The values represent the means and standard deviations from three individual experiments. In all experiments, assayed samples displayed negligible activation of parental HEK-Blue Null2 cells, suggesting specific induction of NF-κB-dependent SEAP expression via the hTLR4/MD-2 signaling pathway in HEK-Blue hTLR4 cells.

As one of the critical mediators induced in response to endotoxin, we assayed the release of TNF-α upon stimulation of human macrophages by lipid IVA samples of KPM318, KPM335 and KPM404. The samples exhibited very low biological activity as demonstrated by their low capacity to provoke TNF-α production in human macrophages even at concentrations of 0.1-1 μg/ml (Figure 7A). Compared to the LPS from the parental strains, which induced maximal TNF-α release at 0.01 μg/ml, TNF-α induction was reduced by about 80-95% even at 100-fold higher levels of mutant extracts. The well-documented ability of lipid IVA to act as an antagonist for the hTLR4/MD-2 signaling pathway [35,36] prompted us to examine the inhibition of the agonistic activity of S-form LPS from Salmonella enterica subspecies enterica serovar Abortusequi (S. Abortusequi) by lipid IVA samples from KPM318, KPM335 and KPM404. As shown in Figure 7B, pre-exposure of macrophages to 0.1 μg/ml and 1 μg/ml of the lipid IVA extracts resulted in 72.0 ± 11.2% and 75.9 ± 2.0% (mean percent inhibition ± SD) inhibition of TNF-α production induced by LPS from S. Abortusequi, respectively. Thus, lipid IVA from KPM318, KPM335 and KPM404 displayed potent antagonistic activity against biologically active wild-type LPS.

Biological activity of LPS from KPM mutants in human macrophages. Macrophages were differentiated from human blood monocytes of healthy donors. On day seven of differentiation, macrophages were seeded at 1 × 10 5 cells/well and stimulated with LPS at the indicated amounts (LPS from S. Abortusequi, BW30270 and BL21 (DE3) at 0.01 μg/ml LPS from strains KPM318, KPM335 and KPM404 at 0.1 μg/ml and 1 μg/ml, respectively) for 4 h at 37°C (A). To determine the antagonistic activity of LPS from KPM strains, macrophages were incubated with LPS samples from KPM318, KPM335 or KPM404 at 0.1 μg/ml or 1 μg/ml for 30 min at 37°C, followed by stimulation of the cells with 0.01 μg/ml of LPS from S. Abortusequi for 4 h (B). Cell-free supernatants were analyzed for TNF-α content by ELISA. The values represent the means and standard deviations from three independent experiments using cells from different donors.

Endotoxin-free expression of ApoA-1 and Hsp70

Previous studies have shown that a BL21 (DE3) ∆lpxM::cat mutant synthesizing a non-myristoylated LPS can be used to express heterologous proteins with reduced stimulatory activity in human LPS-responsive cells [37]. To test the ability of KPM318, KPM335 and KPM404 to serve as hosts for the production of endotoxin-free recombinant proteins, we selected the heterologously expressed human proteins ApoA-1 and Hsp70 as model systems. ApoA-1, a major component of high-density lipoprotein and important mediator in maintaining cholesterol homeostasis [38], is particularly challenging because the 28-kDa protein is known to be directly involved in neutralization of LPS toxicity and, therefore, difficult to separate from endotoxic activity [39,40]. Yet another challenging protein able to associate with LPS is Hsp70 [41]. Moreover, the 70-kDa molecular chaperone has been suggested to function as an endogenous damage-associated molecular pattern protein for activation of the TLR4 signaling pathway upon tissue injury [42]. As Hsp70-induced stimulation of innate immune system cells in many respects resembles the effects of LPS, removal of endotoxin contamination, as frequently present in recombinant Hsp preparations [43], remains a key issue to discriminate between LPS- and Hsp-induced effects.

For production of ApoA-1, we used the T5 promoter-based plasmid pApo404 in E. coli strains BW30270, KPM318 and KPM335, whereas Hsp70 was expressed from pHsp70His under the control of a T7 promoter in BL21 (DE3) and KPM404. SDS-PAGE analysis of the protein samples following minimal purification of each soluble protein extract, by immobilized metal affinity chromatography (IMAC), revealed that the endotoxin-free strains KPM318/pApo404, KPM335/pApo404 and KPM404/pHsp70His produced recombinant ApoA-1 and Hsp70 in approximately equal quantities and had similar impurity profiles as their parental strains, respectively (Figure 8). Since ApoA-1 has been described to also associate with proteins of the host cell [44], it was not surprising to detect relatively high levels of protein contaminants in the IMAC-purified ApoA-1 samples. Regardless, we did not perform any further protein purification or endotoxin removal steps to investigate the biological activity of the ApoA-1 and Hsp70 samples.

SDS-PAGE gels of ApoA-1 and Hsp70. The proteins were expressed in endotoxin-free derivatives of E. coli strains BW30270 (A) and BL21 (DE3) (B), respectively, and minimally purified using IMAC on HisTrap HP (1 ml) columns. The recombinant ApoA-1 and Hsp70 samples (6 μg each) were resolved under denaturing conditions using 12% and 10% polyacrylamide gels, respectively. Molecular mass protein markers (kDa) were run in lanes M. Arrows indicate the positions of ApoA-1 and Hsp70.

The first test of biological activity employed was the Limulus amebocyte lysate (LAL) assay, an FDA-approved method that is based on activation of a coagulation cascade in the LAL by trace amounts of endotoxin [45]. In this assay, the endotoxin unit equivalents determined in ApoA-1 samples from KPM318/pApo404 and KPM335/pApo404 were significantly reduced by 92.7 ± 1.3% and 82.2 ± 3.9% (mean percent inhibition ± SD) compared to those found in the ApoA-1 sample from BW30270/pApo404, respectively (Figure 9). Furthermore, when KPM404/pHsp70His was used as a host for production of Hsp70, the LAL response to the protein decreased by 97.2 ± 0.5% in comparison to the response elicited by Hsp70 obtained from BL21 (DE3)/pHsp70His. The LAL assay, though widely used for the detection and quantitation of endotoxins, is an inappropriate method to discriminate between endotoxically active hexaacylated LPS and endotoxically inactive tetraacylated lipid IVA due to the presence of the LAL-activating 4′-monophosphoryl-diglucosamine backbone in both lipid structures [46,47]. As such, the LAL clotting cascade is activated by a wider spectrum of LPS/lipid A variants than LPS-responsive cells of the human immune system. The residual LAL reactivity of the proteins from the lipid IVA host strains most likely reflects the non-specific nature of the assay, giving rise to false positive endotoxic results.

Reactivity of ApoA-1 and Hsp70 in the Limulus amebocyte lysate (LAL) assay. ApoA-1 was produced in E. coli strain BW30270/pApo404 and its endotoxin-free derivatives KPM318/pApo404 and KPM335/pApo404, whereas Hsp70 was obtained from KPM404/pHsp70His and its parental strain BL21 (DE3)/pHsp70His. The proteins were minimally purified by IMAC and assayed with the LAL test. The measurements represent the means and standard deviations from three individual experiments.

In order to specifically address the endotoxic activity of the ApoA-1 and Hsp70 samples, we utilized the HEK-Blue hTLR4/MD-2 cell activation assay. The ApoA-1 and Hsp70 samples derived from the endotoxin-free strains did not trigger an endotoxic response in HEK-Blue hTLR4 cells, even when present in the assay at 10 μg/ml, whereas the proteins produced in the parental strains showed substantial NF-κB activation already at concentrations in the range between 0.1 μg/ml and 1 μg/ml (Figure 10). These results were in excellent agreement with the inability of KPM318, KPM335 and KPM404 cells and LPS to stimulate the hTLR4/MD-2 signaling pathway (Figures 5 and 6).

Stimulation of hTLR4/MD-2 by ApoA-1 and Hsp70 produced in endotoxin-free E. coli strains. The proteins were minimally purified by IMAC and assayed with HEK-Blue hTLR4 cells for their ability to activate NF-κB-dependent SEAP expression (A and C). HEK-Blue Null2 cells served as a control (B and D). Relative NF-κB induction was measured following stimulation of HEK-Blue hTLR4 and Null2 cells with tenfold serial dilutions of ApoA-1 (A and B) and Hsp70 (C and D) samples obtained by heterologous expression in BW30270/pApo404, KPM318/pApo404 and KPM335/pApo404, and BL21 (DE3)/pHsp70His and KPM404/pHsp70His, respectively. The values represent the means and standard deviations from three individual experiments. In all experiments, the ApoA-1 and Hsp70 samples did not activate NF-κB-dependent SEAP expression in HEK-Blue Null2 cells, suggesting that NF-κB-dependent SEAP expression was due to specific activation of the hTLR4/MD-2 signaling pathway in HEK-Blue hTLR4 cells.

As exemplified herein by heterologous expression of ApoA-1 and Hsp70, proteins prepared from engineered E. coli strains are innately free of endotoxic activity in human LPS-responsive cells, consistent with earlier observations that KPM335 is also a suitable host for production of functional endotoxin-free inclusion bodies of the aggregation-prone fluorescent fusion protein VP1GFP [48]. However, it must be taken into account that lipid IVA may act agonistically in other mammalian hosts such as mouse [49], Chinese hamster [50] or equine [36] cells, which reflects species-specific lipid IVA sensing by the TLR4/MD-2 complex [51,52]. The application of proteins in cells other than human cells may therefore also require lipid IVA depletion. Due to the lack of specificity, the LAL assay cannot be used to assess endotoxic activity but is ideally suited to detect residual lipid IVA in recombinant proteins prepared from LPS-free E. coli strains. We are aware that we currently cannot respond to the question of whether further purification of ApoA1 and Hsp70 would result in complete loss of lipid IVA. However, our data on significantly decreased LAL reactivities of minimally purified ApoA1 and Hsp70 proteins from endotoxin-free bacteria give reason to assume that lipid IVA, being homogeneous, is much easier to remove from downstream products than mature LPS. Despite a common backbone structure, LPS is synthesized as a heterogeneous mixture of structurally related molecules that are decorated with various substituents, usually present in non-stoichiometric amounts [53]. These substitutions, which may vary significantly depending on growth conditions, contribute to considerable physico-chemical heterogeneity of LPS molecules, posing a major challenge to the development of a generally applicable method for endotoxin removal from proteins produced in common E. coli expression strains [5].


RNA Stability and microRNAs

In addition to RBPs that bind to and control (increase or decrease) RNA stability, other elements called microRNAs can bind to the RNA molecule. These microRNAs, or miRNAs, are short RNA molecules that are only 21–24 nucleotides in length. The miRNAs are made in the nucleus as longer pre-miRNAs. These pre-miRNAs are chopped into mature miRNAs by a protein called dicer. Like transcription factors and RBPs, mature miRNAs recognize a specific sequence and bind to the RNA however, miRNAs also associate with a ribonucleoprotein complex called the RNA-induced silencing complex (RISC). RISC binds along with the miRNA to degrade the target mRNA. Together, miRNAs and the RISC complex rapidly destroy the RNA molecule.

Practice Questions

Which of the following are involved in post-transcriptional control?

  1. control of RNA splicing
  2. control of RNA shuttling
  3. control of RNA stability
  4. all of the above

Binding of an RNA binding protein will ________ the stability of the RNA molecule.

  1. increase
  2. decrease
  3. neither increase nor decrease
  4. either increase or decrease

Describe how RBPs can prevent miRNAs from degrading an RNA molecule.

How can external stimuli alter post-transcriptional control of gene expression?

In Summary: Eukaryotic Epigenetic Gene Regulation

In eukaryotic cells, the first stage of gene expression control occurs at the epigenetic level. Epigenetic mechanisms control access to the chromosomal region to allow genes to be turned on or off. These mechanisms control how DNA is packed into the nucleus by regulating how tightly the DNA is wound around histone proteins. The addition or removal of chemical modifications (or flags) to histone proteins or DNA signals to the cell to open or close a chromosomal region. Therefore, eukaryotic cells can control whether a gene is expressed by controlling accessibility to transcription factors and the binding of RNA polymerase to initiate transcription.

To start transcription, general transcription factors, such as TFIID, TFIIH, and others, must first bind to the TATA box and recruit RNA polymerase to that location. The binding of additional regulatory transcription factors to cis-acting elements will either increase or prevent transcription. In addition to promoter sequences, enhancer regions help augment transcription. Enhancers can be upstream, downstream, within a gene itself, or on other chromosomes. Transcription factors bind to enhancer regions to increase or prevent transcription.

Post-transcriptional control can occur at any stage after transcription, including RNA splicing, nuclear shuttling, and RNA stability. Once RNA is transcribed, it must be processed to create a mature RNA that is ready to be translated. This involves the removal of introns that do not code for protein. Spliceosomes bind to the signals that mark the exon/intron border to remove the introns and ligate the exons together. Once this occurs, the RNA is mature and can be translated. RNA is created and spliced in the nucleus, but needs to be transported to the cytoplasm to be translated. RNA is transported to the cytoplasm through the nuclear pore complex. Once the RNA is in the cytoplasm, the length of time it resides there before being degraded, called RNA stability, can also be altered to control the overall amount of protein that is synthesized. The RNA stability can be increased, leading to longer residency time in the cytoplasm, or decreased, leading to shortened time and less protein synthesis. RNA stability is controlled by RNA-binding proteins (RPBs) and microRNAs (miRNAs). These RPBs and miRNAs bind to the 5′ UTR or the 3′ UTR of the RNA to increase or decrease RNA stability. Depending on the RBP, the stability can be increased or decreased significantly however, miRNAs always decrease stability and promote decay.


General Transfection Protocol

Preparing Cells for Transfection

Removing Adherent Cells Using Trypsin

Trypsinizing cells prior to subculturing or cell counting is an important technique for successful cell culture. The following technique works consistently well when passaging cells.

Materials Required:

  • 1X trypsin-EDTA solution
  • 1X PBS or 1X HBSS
  • adherent cells to be subcultured
  • appropriate growth medium (e.g., DMEM) with serum or growth factors or both added
  • culture dishes, flasks or multiwell plates, as needed
  • hemocytometer
  1. Prepare a sterile trypsin-EDTA solution in a calcium- and magnesium-free salt solution such as 1X PBS or 1X HBSS. The 1X solution can be frozen and thawed for future use, but trypsin activity will decline with each freeze-thaw cycle. The trypsin-EDTA solution may be stored for up to 1 month at 4°C.
  2. Remove medium from the tissue culture dish. Add enough PBS or HBSS to cover the cell monolayer: 2ml for a 150mm flask, 1ml for a 100mm plate. Rock the plates to distribute the solution evenly. Remove and repeat the wash. Remove the final wash. Add enough trypsin solution to cover the cell monolayer.
  3. Place plates in a 37°C incubator until cells just begin to detach (usually 1&ndash2 minutes).
  4. Remove the flask from the incubator. Strike the bottom and sides of the culture vessel sharply with the palm of your hand to help dislodge the remaining adherent cells. View the cells under a microscope to check whether all cells have detached from the growth surface. If necessary, cells may be returned to the incubator for an additional 1&ndash2 minutes.
  5. When all cells have detached, add medium containing serum to cells to inactivate the trypsin. Gently pipet cells to break up cell clumps. Cells may be counted using a hemocytometer, distributed to fresh plates for subculturing, or both.

Typically, cells are subcultured to prepare for transfection the next day. The subculture should bring the cells of interest to the desired confluency for transfection. As a general guideline, plate 5 × 10⁴ cells per well in a 24-well plate or 5.5 × 10⁵ cells for a 60mm culture dish for

80% confluency on the day of transfection. Change cell numbers proportionally for different size plates (see Table 3).

Table 3. Area of Culture Plates for Cell Growth.

Size of Plate Growth Area a (cm²) Relative Area b
24-well 1.88 1X
96-well 0.32 0.2X
12-well 3.83 2X
6-well 9.4 5X
35mm 8.0 4.2X
60mm 21 11X
100mm 55 29X

a This information was calculated for Corning® culture dishes.
b Relative area is expressed as a factor of the total growth area of the 24-well plate recommended for optimization studies. To determine the proper plating density, multiply 5 × 10⁴ cells by this factor.

Preparing DNA for Transfection

High-quality DNA free of nucleases, RNA and chemicals is as important for successful transfection as the transfection reagent chosen. See the Protocols and Applications Guide chapter on DNA purification for information about purifying transfection-quality DNA.

Example Protocol Using ViaFect&trade Reagent

We strongly recommend that you optimize transfection conditions for each cell line. If you have optimized transfection parameters, use the empirically determined conditions for your experimental transfections.

If you choose not to optimize transfection parameters, use the general conditions recommended below.

Materials Required:

  • cell culture medium with serum appropriate for the cell type being transfected
  • serum-free cell culture medium for complex formation (such as Opti-MEM® I reduced-serum medium)
  • 96-well or other culture plates
  • U- or V-bottom dilution plates or microcentrifuge tubes

The total volume of transfection complex (medium, DNA and ViaFect&trade Transfection Reagent) to add per well of a 96-well plate is 5&ndash10&mul. The following protocol is a guideline for transfecting approximately 10&ndash20 wells, depending on the volume of ViaFect&trade Transfection Reagent:DNA mixture used. For additional wells, scale volumes accordingly.

  1. To a sterile tube or U- or V-bottom plate, add 90&ndash99&mul of serum-free medium prewarmed to room temperature so that the final volume after adding the DNA is 100&mul. Add 1&mug of plasmid DNA to the medium, and mix. For a 3:1 ViaFect&trade Transfection Reagent:DNA ratio, add 3&mul of ViaFect&trade Transfection Reagent, and mix immediately.
  2. Incubate the ViaFect&trade Transfection Reagent:DNA mixture for 5&ndash20 minutes at room temperature.
    Optional: Add mixture to cells without an incubation period.
    Note: Longer incubations may adversely affect transfections.
  3. Add 5&ndash10&mul of the ViaFect&trade Transfection Reagent:DNA mixture per well to a 96-well plate containing 100&mul of cells in growth medium. We suggest 10&mul of mixture as a starting point. Mix gently by pipetting or using a plate shaker for 10&ndash30 seconds. Return cells to the incubator for 24&ndash48 hours.
    Note: The total growth medium volume may vary depending on well format and your laboratory&rsquos common practices.
  4. Measure transfection efficiency using an assay appropriate for the reporter gene. For transient transfection, cells are typically assayed 24&ndash48 hours after transfection.

Optimizing Transfection with Lipid Reagents

In previous sections, we discussed factors that influence transfection success. Here we present a method to optimize transfection of a particular cell line with a single transfection reagent. For more modern lipid-based reagents such as the ViaFect&trade Transfection Reagent, we recommend initially testing 50&ndash100ng of DNA per well of a 96-well plate at reagent:DNA ratios of 3:1 or 4:1 for adherent cell lines or 2:1 for suspension cell lines. Figure 4 outlines a typical optimization matrix. When preparing the ViaFect&trade Transfection Reagent:DNA complex, the incubation time may require optimization we recommend 5&ndash20 minutes.

Figure 4. Transfection optimization using the ViaFect&trade Transfection Reagent. TF-1 cells were plated in growth media without antibiotics at 30,000 cells per well in a white 96-well assay plate and transfected with a CMV-luc2 plasmid using various lipid (ViaFect&trade Transfection Reagent):DNA ratios. The DNA concentration was held constant at 1&mug per 100&mul of Opti-MEM® I reduced-serum medium, and the amount of ViaFect&trade Transfection Reagent was varied to obtain the indicated ratios. Either 5 or 10&mul of transfection complex was then added to cells in the 96-well plate. Twenty-four hours after transfection, the ONE-Glo&trade + Tox Luciferase Assay was performed. Note that 0:1 is the negative control with DNA but no lipid. These results show that, for this particular cell line, a 2:1 ViaFect&trade Transfection Reagent:DNA ratio gave optimal results.

For traditional cationic lipid reagents, we recommend testing various amounts of transfected DNA (0.25, 0.5, 0.75 and 1µg per well in a 24-well plate) at two charge ratios of lipid reagent to DNA (2:1 and 4:1 see Figure 5). This brief optimization can be performed using a transfection interval of 1 hour under serum-free conditions. One 24-well culture plate per reagent is required for the brief optimization with adherent cells (three replicates per DNA amount).

Figure 5. Suggested plating format for initial optimization of cationic lipid transfection conditions.

A more thorough optimization can be performed to screen additional charge ratios, time points and effects of serum-containing medium at the DNA amounts found to be optimal during initial optimization studies. One hour or two hours for the transfection interval is optimal for many cell lines. In some cases, however, it may be necessary to test charge ratios and transfection intervals outside of these ranges to achieve optimal gene transfer.

Some transfection methods require removing medium with reagent after incubation others do not. Read the technical literature accompanying the selected transfection reagent to learn which method is appropriate for your system. However, if there is excessive cell death during transfection, consider decreasing time of exposure to the transfection reagent, decreasing the amounts of DNA and reagent added to cells, plating additional cells and removing the reagent after the incubation period and adding complete medium.

Endpoint Assays

Many transient expression assays use lytic reporter assays like the Dual-Luciferase® Assay System (Cat.# E1910) and Bright-Glo&trade Assay System (Cat.# E2610) 24 hours after transfection. The Nano-Glo® Dual-Luciferase® Reporter Assay System (Cat.# N1610) allows detection in just a few hours after transfection. However, the time frame for most assays can vary (24&ndash72 hours after transfection), depending on protein expression levels. Reporter-protein assays use colorimetric, radioactive or luminescent methods to measure enzyme activity present in a cell lysate. Some assays (e.g., Luciferase Assay System) require that cells are lysed in a buffer after removing the medium, then mixed with a separate assay reagent to determine luciferase activity. Others are homogeneous assays (e.g., Bright-Glo&trade Assay System) that include the lysis reagent and assay reagent in the same solution and can be added directly to cells in medium. Examine the reporter assay results and determine where the greatest expression (highest reading) occurred. These are the conditions to use with your constructs of interest.

Alternative detection methods include histochemical staining of cells (determining the percentage of cells that are stained in the presence of the reporter gene substrate Figure 6), fluorescence microscopy (Figure 7) or cell sorting if using a fluorescent reporter like the Monster Green® Fluorescent Protein phMGFP Vector (Cat.# E6421).

Figure 6. Histochemical staining of RAW 264.7 cells for &beta-galactosidase activity. RAW 264.7 cells were transfected using 0.1µg DNA per well and a 3:1 ratio of FuGENE® HD to DNA. Complexes were formed for 5 minutes prior to applying 5µl of the complex mixture to 50,000 cells/well in a 96-well plate. Twenty-four hours post-transfection, cells were stained for &beta-galactosidase activity using X-gal. Data courtesy of Fugent, LLC.

Figure 7. Fluorescence microscopy of cells transfected with ViaFect&trade Transfection Reagent. iCell® human tissue cells in 96-well plates were transfected with ViaFect&trade Transfection Reagent and a GFP reporter plasmid at the specified reagent:DNA ratio and GFP expression was imaged after transfection. Panel A. iCell® Hepatocytes with a 6:1 reagent:DNA ratio imaged one day after transfection. Panel B. iCell® Cardiomyocytes with a 2:1 reagent:DNA ratio imaged one day after transfection. Panel C. iCell® DopaNeurons with a 4:1 reagent:DNA ratio imaged three days after transfection. Data courtesy of Cellular Dynamics International.

Real-Time Assays

For some types of transfection experiments, especially those examining the changes in gene expression levels associated with pathological mechanisms, monitoring reporter activity in living cells is desirable. Such real-time assays can provide valuable information on the expression of multiple genes in a dynamic fashion. The Nano-Glo® Live Cell detection options (Cat.# N2011) are designed to detect NanoLuc® luminescence from living cells using nonlytic protocols. These assays monitor luminescence at a single time point or continuously for up to 72 hours without compromising cell viability.


Signaling in cells can occur through protein-protein interactions. Chen et al. describe the design of logic gates that can regulate protein association. The gates were built from small, designed proteins that all have a similar structure but where one module can be designed to interact specifically with another module. Using monomers and covalently connected monomers as inputs and encoding specificity through designed hydrogen-bond networks allowed the construction of two-input or three-input gates based on competitive binding. The modular control elements were used to regulate the association of elements of transcription machinery and split enzymes in vitro and in yeast cells.

The design of modular protein logic for regulating protein function at the posttranscriptional level is a challenge for synthetic biology. Here, we describe the design of two-input AND, OR, NAND, NOR, XNOR, and NOT gates built from de novo–designed proteins. These gates regulate the association of arbitrary protein units ranging from split enzymes to transcriptional machinery in vitro, in yeast and in primary human T cells, where they control the expression of the TIM3 gene related to T cell exhaustion. Designed binding interaction cooperativity, confirmed by native mass spectrometry, makes the gates largely insensitive to stoichiometric imbalances in the inputs, and the modularity of the approach enables ready extension to three-input OR, AND, and disjunctive normal form gates. The modularity and cooperativity of the control elements, coupled with the ability to de novo design an essentially unlimited number of protein components, should enable the design of sophisticated posttranslational control logic over a wide range of biological functions.

Protein-protein interactions are ubiquitous in cellular decision-making and controlling them will be increasingly important in synthetic biology (14). Although protein interactions are central to natural biological circuits, efforts to create new logic circuits have focused on control at the level of DNA (5, 6), transcription (718), or RNA (13, 1922). Recently, protein-based circuits have been generated by rewiring native signaling pathways (2328), bringing proteins together with coiled coils (29), or creating protease cascades (30, 31) however, these circuits were constructed from a limited pool of building blocks, which hinders their scalability. The ability to de novo design protein-based logic gates modulating arbitrary protein-protein interactions could open the door to new protein-based control systems in and out of cells.

In principle, it should be possible to design a wide range of logic gates de novo using a set of heterodimeric molecules. For example, given hypothetical heterodimer pairs A:A′, B:B′, and C:C′, an AND gate modulating the association of A with C′ can be constructed by genetically fusing A′ and B, and B′ and C: association occurs only in the presence of both A′-B, and B′-C (here and below “:” denotes noncovalent interaction and “-” a genetic fusion through flexible linkers). Several building block properties are desirable for constructing such associative logic gates. First, there should be many mutually orthogonal heterodimeric pairs so that gate complexity is not limited by the number of individual elements. Second, the building blocks should be modular and similar in structure so that differences in building block shape and other properties do not have to be considered when constructing the gates. Third, single building blocks should be able to bind to multiple partners with different and tunable affinities, allowing inputs to perform negation operations by disrupting preexisting lower-affinity interactions. Fourth, the interactions should be cooperative so that gate activation is not sensitive to stoichiometric imbalances in the inputs. In the above AND gate, for example, if the interactions are not cooperative, then a large excess of A′-B will pull the equilibrium toward partially assembled complexes (A′-B with either A or B′-C but not both), which will limit gate activation.

Here, we explored the possibility of designing logic gates satisfying all four of the above criteria using de novo–designed protein heterodimers with hydrogen bond network–mediated specificity (32). Sets of mutually orthogonal designed heterodimers (DHDs, hereafter referred to by numbers, e.g., 1 and 1′ form one cognate pair table S1) with hydrogen bond network–mediated specificity (e.g., see Fig. 1A, inset) are available for logic gate construction, satisfying condition 1 (orthogonality). The heterodimeric interfaces all share the same four helix bundle topology (Fig. 1A), satisfying condition 2 (modularity). The shared interaction interface allows a limited amount of cross-talk between pairs, leading to a hierarchy of binding affinities, satisfying condition 3 (multiple binding specificities). Inspired by cooperative systems in nature (33, 34), we sought to achieve condition 4 (cooperativity) by constructing the monomer fusions (A′-B and B′-C in the above example) in such a way that the interaction surfaces (with A and C′) are buried within the fusions. The free energy required to expose these buried interfaces would oppose gate activation, and we reasoned that the system could be tuned so that the sum of the binding energies of the two partners, but not either one alone, would be sufficient to overcome this barrier, ensuring cooperative gate activation. If condition 2 (modularity) holds, then a single scheme for ensuring cooperativity could in principle work for a wide range of gate configurations.

(A) Left: Backbone structure of A:A′ heterodimer building block, with its hydrogen bond network shown in the inset. Bottom: Shorthand representations used throughout figures. (B) Thermodynamic cycle describing the induced dimerization system. (C) Simulation of the induced dimerization system under thermodynamic equilibrium. A and B′ monomers were held constant at 10 μM each while titrating in various initial amounts of the A′-B dimerizer proteins. If binding is not cooperative (small c), then the final amount of trimeric complexes decreases when the dimerizer protein is in excess. (D) Equilibrium denaturation experiments monitored by CD for designs with 6- and 12-amino acid (AA) linkers. Circles represent experimental data, and lines are fits to the three-state unimolecular unfolding model. (E) Experimental SAXS profile of 1′-2′ with a six-residue linker (in black) fitted to the calculated profile of 1:1′ heterodimer (in red). (F) Schematic of induced dimerization system (with a six-residue linker) experimental results in (G) and (H). (G) nMS titration of 2 against 10 μM of 1′-2′ in the presence (red) or absence (blue) of 10 μM of 1. (H) nMS titration of 1′-2′ against 10 μM each of 1 and 2. Dimer 1 and 2 refer to partial dimeric complexes consisting of the dimerizer binding to either of the monomers. For comparison, the thermodynamic model result with c = 991,000 is shown in cyan. (I) Schematic of testing of the induced dimerization system in yeast, with in vivo results shown in (J). Pg, progesterone. (K) Two-input AND gate schematic, with nMS titration results shown in (L). Trimer 1 and 2 refer to partial trimeric complexes of the two dimerizer proteins binding to either one of the monomers. (M) A three-input AND gate, with nMS titration results shown in (N). Tetramer 1 and 2 refer to partial tetrameric complexes of the three dimerizer proteins binding to either one of the monomers. All error bars are standard deviations of n = 3 independent replicates.

To explore the design of cooperative building blocks, we focused on the simple system A + A′-B + B′ (we refer to this as induced dimerization below, A and B′ as the monomers, and A′-B as the dimerizer). If binding is not cooperative, then the amount of the trimeric complex decreases when A′-B is in stoichiometric excess relative to A and B′: the formation of intermediate dimeric species of the dimerizer binding to either of the monomers competes with formation of trimeric complexes. On the contrary, if binding is cooperative such that no binding to either monomer occurs in the absence of the other, then the amount of trimeric complex formed becomes insensitive to an excess of the dimerizer. A simple thermodynamic model of the effect of binding cooperativity on the stoichiometry dependence of such induced dimerization systems (Fig. 1B and see modeling section in the supplementary materials) shows that, as the binding cooperativity decreases, there is a corresponding decrease in the population of full trimeric complexes at high dimerizer concentrations (Fig. 1C).

We hypothesized that a folded four-helix bundle–like state of the A′-B dimerizer could oppose binding to either A or B′ because the relatively hydrophobic interacting surfaces would likely be sequestered within the folded structure (fig. S1A). We tested different flexible linker lengths connecting A′ with B using heterodimers 1:1′ and 2:2′ as a model system. At all linker lengths tested (between 0 and 24 residues), constructs were folded and stable in circular dichroism (CD) guanidine hydrochloride denaturation experiments, with unfolding free energies >13 kcal/mol (Fig. 1D, fig. S2, and table S3). Although 1′-2′ dimerizer constructs with short linkers of 0 and 2 residues, or with a very long 24-residue linker, could be purified as monomers (fig. S1B), they were prone to aggregation, perhaps due to domain swapping. By contrast, designs with 6 and 12 residue linkers remained largely monomeric (table S4). Small-angle x-ray scattering (SAXS) experiments (35) indicated that their hydrodynamic radii are close to those of folded four-helix bundle DHDs (Fig. 1E and table S2). Linkers in this length range likely allow the two monomers (1′ and 2′) to fold back on each other such that the largely hydrophobic interaction surfaces are buried against each other such a structure would have to partially unfold for 1′-2′ to interact with either 1 or 2. The magnitude of the unfolding energy (ΔGopen in Fig. 1B) determines the extent of cooperativity for the gate. We selected linker lengths of 6, 10, or 12 residues for all of the following experiments.

We studied the cooperativity of the induced dimerizer system in vitro using native mass spectrometry (nMS, 36, 37), which can directly measure the populations of different oligomeric species in a sample (tables S5 to S8 for calibration curve, see fig. S3). We first measured the extent to which 1 activates the binding of 2 to 1′-2′ (Fig. 1F). 1, 2, and 1′-2′ were separately expressed in Escherichia coli and purified. At 10 μM of 1′-2′ and 20 μM of 2, we observed a 33 fold increase in binding between 2 and 1′-2′ upon addition of 10 μM of 1 (Fig. 1G), a fold increase comparable to naturally occurring allosteric systems (33). To assess the sensitivity of binding to stoichiometric imbalance, 10 μM 1 and 2 were titrated with increasing concentrations of 1′-2′ (Fig. 1H) and the species formed were determined by nMS. The heterotrimeric 1:1′-2′:2 complex was observed over a wide range of 1′-2′ concentrations (Fig. 1H). Even in the presence of a 6-fold excess of 1′-2′, there was no decrease in the amount of 1:1′-2′:2 formed, and neither 1:1′-2′ nor 1′-2′:2 was detected (Fig. 1H). We define a cooperativity parameter c as the ratio of the affinities in the presence and absence of the other monomer, which in our model directly relates to the free energy of opening of the dimerizer ( c = e Δ G o p e n / RT see supplementary materials). The estimated c value from fits of the thermodynamic model to nMS data (Fig. 1H, cyan line) is 991,000 ± 21 (for reference, the c value of the naturally occurring N-Wasp system is 350 but system differences complicate quantitative comparisons). This value of c corresponds to ΔGopen of 8.2 kcal/mol, which is about half the measured unfolding free energy of 1′-2′ (table S3), suggesting that binding may not require complete unfolding of the four-helix bundle state of the dimerizer.

To investigate the cooperativity of the induced dimerizer system in living cells, we used a two-hybrid–like assay in yeast. 11′ was fused to the DNA-binding domain ZF43 (14), 7 to the transactivation domain VP16, and the dimerizer 11-7′ was placed under the control of a progesterone-responsive element. Association of the DNA-binding and activation domains results in transcription of red fluorescent protein (RFP) (Fig. 1I). Treating cells with increasing amount of progesterone resulted in up to a 4.5-fold increase in RFP signal, with only a small drop at saturating progesterone concentrations (Fig. 1J). On the basis of calibration curves, under these conditions, 11-7′ is expected to be in >5-fold molar excess over 11′ and 7 (fig. S4), suggesting that 11-7′ binds cooperatively to 11′ and 7 in cells. Thus, the cooperativity of the dimerizer system makes it robust to fluctuating component stoichiometries in cells.

With dimerizers displaying cooperative binding, we reasoned that the lack of dependence on stoichiometric excesses of one of the components should extend to more complex gates. Using nMS, we investigated the cooperativity of a two-input AND gate constructed with the two dimerizers 1′-3′ and 3-2′ as inputs and monomers 1 and 2 brought together by the two inputs (Fig. 1K). As the concentration of the 2 inputs increased, the amount of heterotetrameric complex plateaued at a stoichiometry of 2:1, and then largely remained constant with a small drop at molar ratio of 6:1. Only very small amounts of partial complexes (heterotrimers and heterodimers) were observed, further indicating high cooperativity (Fig. 1L). We constructed a three-input AND gate from 1′-4′, 4-3′, and 3-2′, which together control the association of 1 and 2 (Fig. 1M). Similar to the two-input AND gate, the abundance of full, pentameric complexes only decreased slightly at greater than stoichiometric concentrations of inputs with no detectable competing tetrameric complexes (Fig. 1N).

We explored the modular combination of DHDs (table S1) to generate a range of two-input cooperatively inducible protein heterodimer (CIPHR) logic gates. Monomers from individual DHDs were linked to effector proteins of interest by genetic fusion such that the inputs (linked heterodimer subunits) control colocalization or dissociation of the effector proteins. Taking advantage of previously measured all-by-all specificity matrices for the DHDs (32), we explored constructing gates from two interaction modalities: cognate binding between designed protein pairs and competitive binding involving multispecific interactions (Fig. 2A).

(A) CIPHR gates are built from DHDs (top) with monomers or covalently connected monomers as inputs (left) some gates use only the designed cognate interactions (left side of middle panel), whereas others take advantage of observed binding affinity hierarchies (right side of middle panel). (B and C) Two-input AND (B) and OR (C) CIPHR logic gates based on orthogonal DHD interactions. (D to G) NOT (D), NOR (E), XNOR (F), and NAND (G) CIPHR logic gates made from multispecific and competitive protein binding. For each gate, black dots represent individual Y2H growth measurement corrected over background growth, with their average values shown in green bars. Components in gray boxes indicate the DHD pairs used. Blue boxes indicate affinity gradients. *No yeast growth over background. “0” and “1” in the middle and right blocks represent different input states and expected outputs, respectively.

We began by constructing AND and OR gates, reading out gate function using a yeast-two-hybrid (Y2H) setup similar to previously described yeast-four-hybrid systems (38, 39). To construct an AND gate, we fused 2 to the Gal4 activation domain (AD) and 1 to the Gal4 DNA-binding domain (DBD). In this scheme, the colocalization of AD and DBD, and the resulting transcriptional activation of the His3 gene, should require the expression of both input proteins (1′-5, 5′-2′). Indeed, growth in medium lacking histidine required expression of both inputs (Fig. 2B). An OR gate was similarly constructed by linking the 1-6 fusion to the AD and 7′ to the DBD. Expression of either of the inputs, 1′-7 or 6′-7, resulted in growth by driving association of AD with DBD (Fig. 2C).

We explored the construction of additional Boolean logic gates by exploiting binding affinity hierarchies identified in all-by-all Y2H experiments (32). 8 alone formed a homodimer, but in the presence of 8′ it dissociated to form the 8:8′ heterodimer (fig. S5A). We constructed a NOT gate by fusing 8 to both AD and DBD the 8:8 homodimer supported yeast growth but in the presence of coexpressed 8′ input protein, the interaction was broken and growth was slowed (Fig. 2D). On the basis of the affinity hierarchy 9:9′10:10′ > 9:10′ (fig. S5B), we constructed a NOR gate in which 9 was fused to the AD and 10′ to the DBD, with 9′ and 10 as the two inputs. Either or both inputs outcompeted the 9:10′ interaction and hindered yeast growth (Fig. 2E). On the basis of the affinity hierarchy 9′:1′ > 9:9′1:1′ > 9:1 (fig. S5B), an XNOR gate was constructed by fusing 9 to AD, 1 to DBD, and using 9′ and 1′ as the two inputs: the presence of either outcompeted the 9:1 binding and blocked growth, but when both were expressed they instead interacted with each other and growth was observed (Fig. 2F). Similarly, a NAND gate was designed based on the interaction hierarchy 1′:10′ > 1:1′10:10′ (fig. S5B). Neither 1 nor 10 alone could outcompete the 1′:10′ interaction and hence growth occured, but when both were expressed, the free energy of formation of both 1:1′ and 10:10′ outweighed that of 1′:10′ and growth was blocked (Fig. 2G).

We next investigated three-input CIPHR logic gates. We first used nMS to characterize a three-input AND gate (Fig. 1M) in which monomers 1 and 2 are brought into proximity by the three inputs 1′-4′, 4-3′, and 3-2′. We experimentally tested all eight possible input combinations (Fig. 3A) with both 1 and 2 present, quantifying all complexes using nMS. Consistent with three-input AND gate function, 1 and 2 only showed significant coassembly when all three inputs were present (Fig. 3B).

(A) Schematic of a three-input AND gate. (B) nMS results indicating proper activation of the three-input AND gate only in the presence of all three inputs. (C) Schematic of a three-input OR gate. (D) Y2H results confirming activation of the three-input OR gate with any of the inputs. (E) Schematic of a DNF gate. (F) Y2H results confirming proper activation of the gate. For each gate, black dots represent individual measurements with their average values shown in green bars. For Y2H-based measurements [(D) and (F)], the growth measurements are corrected over background growth. Components in gray boxes indicate the DHD pairs used.

To test three-input CIPHR gate function in cells, we designed two additional gates using the same four pairs of DHDs and tested them by Y2H. To make a three-input OR gate, 1′-6-7 was fused to AD and 11′ to DBD. Any one of the three inputs (11-1, 11-6′, or 11-7′) connects the AD to the DBD through 1′, 6, or 7, respectively (Fig. 3C). Y2H results confirmed the expected behavior of this logic gate in cells: any of the input proteins induced cell growth (Fig. 3D). We additionally constructed a CIPHR-disjunctive normal form [DNF (A AND B) OR C] gate by fusing 1′-6 to AD and 11′ to DBD with inputs 11-7′, 7-1, or 11-6′ (Fig. 3E). In Y2H experiments, the DNF gate functioned as designed, with low yeast growth levels when no input or only one of the 11-7′ and 7-1 input proteins was present and high yeast growth levels otherwise (Fig. 3F).

To test the transferability of CIPHR logic gates, we explored the ability of CIPHR logic gates to reconstitute split enzyme activity by controlling the association of the two halves of the NanoBiT split luciferase system (4042). Monomers from 1:1′, 2:2′, 4:4′, and 9:9′ (Fig. 4A) were fused in pairs to the two split domains (smBiT and lgBiT) and produced by in vitro transcription and translation, which facilitated a rapid testing cycle enabling the full 4×4 interaction affinity hierarchy to be determined by monitoring luciferase activity after mixing (fig. S6A). On the basis of this hierarchy, we constructed and experimentally verified an induced dimerization circuit with 4-smBiT, 1-lgBiT, and 1′-4′ as the input (Fig. 4B and fig. S6, C and D) characterization of the time dependence of the response revealed a 7-fold increase in signal 5 min after adding inputs (fig. S6D). We also constructed an AND gate with 4-smBiT, 1-lgBiT, and 1′-2 and 2′-4′ as the inputs (Fig. 4C) and a NOR gate with 1′-smBiT, 2′-lgBiT, and 1 and 2 as the inputs (Fig. 4D), both of which had the designed dependence of gate function (i.e., luciferase activity) on the inputs. We investigated the response of the NOR gate to varying concentrations of the inputs against the NanoBiT components held at 5 nM and found a sharp drop in signal around 5 nM for both inputs, consistent with NOR logic (Fig. 4E and fig. S6E).

(A) Four pairs of DHDs were modularly combined to construct CIPHR logic gates that can be used to control different functions: catalytic activity of split luciferase (B to E) and gene expression in primary human T cells (F and G). (B) the induced dimerization system, (C) AND gate, and (D) NOR gate coupled to NanoBiT split luciferase system, tested by in vitro translation and monitoring luminescence. (E) In vitro titration of the two inputs of the NOR gate in D while keeping 1′-smBiT and 2′-lgBiT fixed at 5 nM. NOT gate (F) and OR gate (G) using a split TALE-KRAB repression system to control expression of TIM3 proteins in primary human T cells, tested by flow cytometry.

Engineered T cell therapies are promising therapeutic modalities (4345) but their efficacy for treating solid tumors is limited at least in part by T cell exhaustion (46, 47). Immune checkpoint genes including TIM3 are believed to play critical roles in modulating T cell exhaustion (4850). To put the transcription of such proteins under the control of the CIPHR logic gates, we took advantage of potent and selective transcriptional repressors of immune checkpoint genes in primary T cells that combine sequence-specific transcription activator-like effector (TALE) DNA-binding domains with the Krüppel-associated box (KRAB) repressor domain (51). Repression activity is preserved in split systems pairing a DNA recognition domain fused to a DHD monomer with a repressor domain fused to the complementary DHD monomer (51). We reasoned that this system could be exploited to engineer programmable therapeutic devices by making the joining of the DNA recognition and transcriptional repression functionalities dependent on CIPHR gates. Use of a repressive domain effectively reverses the logic of CIPHR gates when expression level of the target gene is measured as the output.

To test the feasibility of this concept, we used a TALE-KRAB fusion engineered to repress the immune checkpoint gene TIM3 (51). We designed a NOT gate with 1 fused to the TALE DNA recognition domain, 9′ fused to KRAB, and the 1′-9 dimerizer protein as the input (see fig. S7A for T cell DHD specificity matrix). In this scheme, 1′-9 brings KRAB to the promoter region bound by the TALE, therefore triggering repression of TIM3 (Fig. 4F). Taking advantage of the interaction between 9 and 1′, we built an OR gate with 9-TALE and 1′-KRAB fusions TIM3 was repressed in the absence of inputs but upon addition of either 9′ or 1, the weaker 9:1′ interaction was outcompeted in favor of the stronger 9:9′ and 1:1′ interactions, restoring TIM3 expression (Fig. 4G). These results suggest that the combination of CIPHR and TALE-KRAB systems could be directly applied to add signal-processing capabilities to adoptive T cell therapy.

The systematic design of logic gates described herein takes advantage of the strengths of de novo protein design. Because the building block heterodimers are designed de novo, many more components for gate construction with nearly identical overall topology can be generated than are available by repurposing biological motifs. The encoding of specificity using designed hydrogen bond networks enables a wide range of binding affinities between monomers with similar structures, which in turn allows the construction of more complex gates that are based on competitive binding. From the protein biophysics perspective, our results highlight the strong synergy between de novo design of protein complexes and nMS and, more generally, the ability of de novo protein design to generate complex cooperative assemblies. For example, detecting and quantifying the 33-fold activation of binding in Fig. 1G depended critically on the ability to resolve all species formed in solution by nMS. Analysis of the three-input logic gates in Fig. 3B required distinguishing the designed heteropentameric assemblies, which are composed of five distinct protein chains, from the very large number of alternative possible heterotetrameric, trimeric, and dimeric complexes. The ability to generate highly cooperative and well-defined assemblies composed of five distinct polypeptide chains demonstrates that de novo protein design is starting to approach the complexity of naturally occurring protein assemblies, which are responsible for much of biological function.

Unlike nucleic acid–based logic gates, CIPHR gates can be directly coupled to arbitrary protein actuation domains, offering greater diversity in the types of functional outputs. We illustrate here the coupling to transcriptional activation and repression and split enzyme reconstitution in principle, any function that can be modulated by protein-protein association can be put under the control of the CIPHR gates. Because the designed components are hyperstable proteins and no additional cellular machinery is required, the gates should function in a wide range of conditions inside and outside of cells (here, we have demonstrated function with purified components in cell-free extracts, yeast cells, and T cells). The small size of DHDs and thus their genetic payload makes them attractive for mammalian cell engineering. The sophistication of the circuits could be further increased by proteolytic activation as in recent elegant studies using protease-based protein circuits (30, 31) our purely protein interaction–based circuits have advantages in bioorthogonality, demonstrated scalability to three inputs, composability (the output, like the input and the computing machinery, consists of interactions between building blocks with common design features), and extensibility because an essentially unlimited repertoire of heterodimeric building blocks can be created using de novo design.


Enzymes

At the most basic definition, enzymes are specialized proteins that initiate changes in the body. In the trillions of biochemical reactions happening in our cells every minute, enzymes play the key role of catalyzing those reactions. This is not to say that those reactions would not occur normally, but enzymes speed up the rate of those processes. Enzymes are reusable proteins that are tailored for a specific type of reaction within a series or cycle of reactions. A substrate is turned into a product through its interaction with an enzyme. That product will often become the substrate for the next reaction in a metabolic cycle or pathway, which will require a different type of enzyme.

The delicate interplay of enzymes and elaborate metabolic pathways is what sustains critical processes we need to survive, from the generation of usable energy to the replication of DNA. There are approximately 3,000 different enzymes found within the body, each serving a unique and valuable purpose to our cells, tissues and organs!


Research could enable biotechnology advances: Medicine, protective equipment, sensors

Credit: The Army Research Laboratory

New Army-funded synthetic biology research manipulated micro-compartments in cells, potentially enabling bio-manufacturing advances for medicine, protective equipment and engineering applications.

Bad bacteria can survive in extremely hostile environments—including inside the highly acidic human stomach—thanks to their ability to sequester toxins into tiny compartments.

In a new study, published in ACS Central Science, Northwestern University researchers controlled protein assembly and built these micro-compartments into different shapes and sizes, including long tubes and polyhedrons. Because this work illuminates how biological units, such as viruses and organelles, develop, it also could inform new ways to design medicine, synthetic cells and nano-reactors that are essential for nanotechnology.

"These results are an exciting step forward in our ability to design complex protein-based compartments," said Dr. Stephanie McElhinny, program manager at the U.S. Army Combat Capabilities Development Command, known as DEVCOM, Army Research Laboratory. "Being able to control the size and shape of these compartments could enable sophisticated bio-manufacturing schemes that are customized to support efficient production of complex molecules and multi-functional materials that could provide the future Army with enhanced uniforms, protective equipment and environmental sensors."

Further down the road, these insights potentially could lead to new antibiotics that target micro-compartments of pathogens while sparing good bacteria.

Researchers control protein assembly and build cell micro-compartments into different shapes and sizes that could lead to bio-inspired building blocks for various engineering applications.

"By carefully designing proteins to have specific mutations, we were able to control assembly of the proteins that form bacterial micro-compartments," said Dr. Monica Olvera de la Cruz, professor of materials science and engineering and chemistry at Northwestern who led the theoretical computation. "We used this also to predict other possible formations that have not yet been observed in nature."

Researchers control protein assembly and build cell microcompartments into different shapes and sizes that could lead to bio-inspired building blocks for various engineering applications Credit: Monica Olvera de la Cruz, Northwestern University

Many cells use compartmentalization to ensure that various biochemical processes can occur simultaneously without interfering with one another. Made of proteins, these micro-compartments are a key to survival for a wide variety of bacterial species.

"Based on previous observations, we have known that the geometry of micro-compartments can be altered," said Dr. Danielle Tullman-Ercek, associate professor of chemical and biological engineering at Northwestern who led the experimental work. "But our work provides the first clues into how to alter them to achieve specific shapes and sizes."

To study these crucial compartments, the Northwestern team turned to Salmonella enterica, which rely on micro-compartments to break down the waste products of good bacteria in the gut. When the researchers genetically manipulated a protein isolated from Salmonella, they noticed the micro-compartments formed long tubes.

"We saw these weird, extended structures," Tullman-Ercek said. "It looked like they used the varying building blocks to form different shapes with different properties."

By coupling the mechanical properties of the compartment with the chemicals inside the compartment, Olvera de la Cruz and her team used theoretical computation to predict how different mutations led to different shapes and sizes. When six-sided proteins assembled together, they formed long tubes. When five-sided proteins assembled together, they formed soccer ball-shaped icosahedrons. The team also predicted that proteins could assemble into a triangular samosa shape, resembling the fried, South Asian snack.

Understanding this process could lead to bio-inspired building blocks for various engineering applications that require components of varying shapes and sizes.

"It's like building with Legos," Tullman-Ercek said. "It's not desirable to use the same shape block over and over again we need different shapes. Learning from bacteria can help us build new and better structures at this microscopic scale."


The Institute for Creation Research

The March 1998 Impact article"Cloning - What is It and Where is It Taking Us?" discussed the procedure of cloning by somatic cell transfer. In that procedure, the nucleus from a cell derived from an embryo, a fetus, or tissue of an adult is inserted into an egg from which the nucleus has been removed. After the egg develops to the appropriate stage, the embryo is inserted into the uterus of a properly prepared female and allowed to develop to term. This produces an offspring essentially genetically identical to the animal that provided the nucleus that was inserted into the egg. This article will discuss the transfer of genes from one species to another in order to endow the recipient species with beneficial properties, or to enable the recipient to produce human proteins for injection into human patients who lack a vitally important protein.

Production of Therapeutic Proteins by Gene Transfer

There are many proteins essential to good health that some people cannot produce because of genetic defects. These proteins include various blood-clotting factors causing hemophilia, insulin (resulting in diabetes), growth hormone (resulting in lack of proper growth), and other proteins, the administration of which corrects pathological conditions or results in other therapeutic benefits. The early work in this field employed bacteria. Some bacteria in a bacterial culture may contain small circular DNA molecules called plasmids. These plasmids are not part of the chromosomal DNA that is possessed by all the bacteria of the culture. As these exceptional bacterial cells reproduce by binary fission or cell division, the plasmids are transmitted to the daughter cells. They can also be transmitted to other cells by conjugation. Scientists have learned how to utilize plasmids to transfer human genes to bacterial cells. If the gene inserted into the plasmid of bacteria is the human gene for insulin, for example, the bacteria into which this gene is inserted produces human insulin.

Insertion of a DNA section into a plasmid

Scientists have identified and isolated enzymes (called restriction enzymes, or restriction endonucleases) each of which cuts genes in very specific places. More than 100 of these enzymes have been isolated. After the position of the human gene that codes for the desired therapeutic protein has been located on the chromosome, the gene is cut out using the appropriate restriction enzymes and the gene is isolated. The same restriction enzymes are used to cut out a piece of the circular DNA plasmid. Thus, the two ends of the human gene will be those that will link up with the open ends of the plasmid. An enzyme called DNA ligase is used to couple each end of the gene to the open ends of the plasmid, restoring a circular DNA molecule with the human gene replacing the piece cut out of the plasmid. These plasmids, now including the human gene, are reinserted into bacteria. These bacteria are cultured, producing large quantities of identical bacteria carrying the human gene that is reproduced along with the bacterial DNA. Furthermore, these bacteria produce the human protein coded for by the spliced human gene. The protein is isolated from the bacterial culture, purified, and injected into those patients suffering from pathological conditions because their bodies cannot produce sufficient quantities of the protein.

Before genetic engineering, these proteins had to be painstakingly isolated from tissues or blood, but since they are produced in such minute quantities, the isolation of significant quantities required the processing of large quantities of material. As a result, they were very expensive. Relatively much larger quantities were obtained from genetically altered bacterial cultures, but the cost, although less, was still high. The bioreactors (that is, the machinery required to culture large quantities of the bacteria containing the human gene) are enormously expensive and must be operated by several scientists and technical assistants. Furthermore, the proper operation of the bioreactor is sensitive to small changes in temperature and composition of the culture broth.

Fortunately, an alternative method has been developed which promises to be much less expensive and much more efficient. This method utilizes an animal, such as a pig, as the bioreactor. It required years for scientists to design, develop, and build the very expensive, difficult to operate bioreactor devised by man. God had already devised a much more efficient, much cheaper bioreactor. Scientists finally realized that it would be possible by genetic manipulation to induce a female pig, cow, or other animal to produce the desired human proteins in its milk.

Genetic engineering of a milk protein

This procedure has now been successful in both pigs and cows. Among animals, the pig has a number of advantages. Its gestation period is only four months. At 12 months of age the pig is fertile and produces large litter sizes (usually 10 to 12 piglets). A lactating pig produces 300 liters (about 315 quarts) of milk in a year. The procedure is carried out as follows:

  1. The DNA fragment (gene) that codes for the needed human protein is isolated.
  2. The DNA fragment that promotes production of proteins in mammary glands is isolated and linked or combined to the human gene.
  3. Fertilized eggs from a donor pig are obtained.
  4. The human DNA is injected into an egg in the region of the male pronucleus (DNA from the sperm before it unites with DNA of the egg) using a very slim micro pipette. The human DNA is thus incorporated into the pig nuclear DNA.
  5. The egg is implanted in the uterus of another pig and develops into a newborn female pig.
  6. The desired protein is isolated from the milk of the female pig once grown.

This procedure was carried out successfully by American scientists and the results were published in 1994. 1 The human gene they used codes for Protein C that acts to control blood clotting. They obtained one gram of Protein C from each liter of milk produced by the pig. This is 200 times the concentration found in human blood plasma. Only about one third of the Protein C obtained was biologically active. The reason for this is that many modifications of a protein must be performed in a cell after the protein chain is formed. For example, sections are cut out of the protein complex sugar groups may be attached at certain places in the protein chain and cell wall anchor groups may be added. The scientists discovered that a key processing enzyme, called furin, was present in insufficient quantities, so they added to their gene complex the gene that codes for furin. This increased the yield of active Protein C. The human proteins produced in this way must be tested for safety and effectiveness. At this writing, an anti-clotting protein called anti-thrombin is now being tested in clinical trials.

Comparing this procedure to the use of bioreactor machines illustrates the fact that a generation of biochemical engineers failed to match the abilities of a tool for making proteins (the pig) that God had prepared. The mammary gland is optimized to maintain a high density of cells to deliver to them an ample supply of nutrients and to channel proteins that are produced in a form that can easily be isolated and purified. This procedure has proven to be successful and promises to provide a method for producing valuable therapeutic proteins at much lower cost.