Analysis of Cationic-Lipid Plasmid-DNA Complexes
`
`7 2 4 0 A N A LY T I C A L C H E M I S T R Y / O C T O B E R 1 , 2 0 0 7
`
`© 2 0 0 7 A M E R I C A N C H E M I C A L S O C I E T Y
`
`See https://pubs.acs.org/sharingguidelines for options on how to legitimately share published articles.
`
`Downloaded via PURDUE UNIV on November 8, 2018 at 23:23:16 (UTC).
`
`PROTIVA - EXHIBIT 2012
`Moderna Therapeutics, Inc. v. Protiva Biotherapeautics, Inc.
`IPR2018-00739
`
`

`

`Analysis of Cationic-Lipid Plasmid-DNA Complexes
`
`Complexes between plasmid
`
`DNA and various cationic
`
`lipids are promising
`
`vehicles to deliver
`
`genetic information
`
`into cells for gene
`
`therapy or vaccines.
`
`C. Russell Middaugh
`Joshua D. Ramsey
`
`University of Kansas The idea of using genes as drugs
`
`is both exciting and controver-
`sial. In most gene therapies, the proteins that are coded
`for by the DNA have a therapeutic effect, but the genetic material itself
`is delivered to the patient, generally in two different ways. The first uses
`modified viruses that contain the gene(s) of interest. This takes advan-
`tage of the fact that viruses have evolved over hundreds of millions of
`years to efficiently deliver their genomes into cells and have their content
`expressed. This can work fairly well, but the problems of immunogenic-
`ity and toxicity, even to the extent of death, have led to serious concerns
`about this approach (1, 2).
`The second method attempts to build a synthetic virus—an approach
`that provides better control of the delivery system’s properties. These
`nonviral vectors usually are created by combining a bacterial plasmid
`containing the gene(s) of interest with a positively charged (usually
`polymeric) partner to form a molecular complex. Plasmids themselves are
`manipulated easily to contain any gene of interest as well as on-and-off
`elements and other regulatory signals. Preparation of large quantities in
`a pure form is also straightforward. Positively charged complexing agents
`that have been used so far include basic peptides, polyethylenimine,
`amino dendrimers, various synthetic block copolymers, and cationic lip-
`ids (CLs) (3–7). Anionic lipids in combination with multivalent cations
`have been explored as an alternative to CLs (8). The CL-containing
`vectors are, however, the most thoroughly investigated. They have been
`shown to have some efficiency in the clinic, although it is significantly
`less than that of their viral counterparts; cytotoxicity and immunogenic-
`ity are still problematic (9, 10). Reviews have addressed characterization
`and the biological aspects of CLs for gene delivery (11–13).
`Traditionally, gene delivery vectors have been analyzed in terms of
`their biological effects, such as their ability to enter cells and direct the
`expression of selected encoded genes. This process is known as trans-
`fection, and its measurement lacks the accuracy and precision expected
`for the analysis of a material that will be used as a drug in humans.
`Furthermore, the complexity of biological fluids makes physical char-
`acterization of vectors in such environments difficult. Characterization
`
`O C T O B E R 1 , 2 0 0 7 / A N A LY T I C A L C H E M I S T R Y 7 2 4 1
`
`

`

`of vectors in situ typically requires the vector to be
`radio- or fluorescently labeled. Thus, measurements
`are performed most often on pure, buffered solutions of
`DNA–polycation complexes. The vectors themselves are large,
`complex, and usually very heterogeneous in structure. This
`makes their physical and chemical characterization difficult,
`even in a pure state.
`Recent attempts have been made to treat such vectors with
`rigorous physicochemical methods to produce stabler, more
`effective, and safer gene-based drugs. In this article, we take
`CL–DNA complexes (CLDCs), or “lipoplexes”, as proto-
`typical examples and discuss the progress and
`future of their physical and chemical analysis.
`Originally described in 1987 (7), CLDCs are
`the most thoroughly studied nonviral gene de-
`livery agents and serve as a good introduction
`to the analytical problems involved in their
`characterization.
`
`ject to a variety of chemical changes. The simplest
`approach is to separate the DNA from the CL by
`methods based on size, charge, or density. This can
`work only if the complexes can be dissociated quantitatively.
`Because the interactions between the CL and DNA are pri-
`marily electrostatic in nature, in principle, separation can
`be achieved by adding salt, but the presence of additional,
`less clearly defined interactions makes this difficult if not
`impossible.
`If separation can be accomplished, analysis of the individual
`components is easy with conventional methods. For example,
`
`(I) C0 < O
`(II) (cid:1)
`
`Primary structure and composition
`When highly negatively charged DNA is mixed
`with virtually any polycation, some degree of
`condensation of the polynucleotide occurs. In
`solution, plasmid DNA usually exists as a
`highly supercoiled, covalently closed circle
`of DNA with small amounts of open circular
`(nonsupercoiled) and linear (cleaved in both
`polynucleotide chains) contaminants. If you
`take a rubber band and twist it several times,
`you will observe its collapse into a much smaller
`volume. This is essentially what happens when
`the negative charges on the phosphate groups
`of the DNA are neutralized by polycations,
`which relieve the electrostatic repulsive forces
`within the DNA and subsequently reduce its
`volume.
`In the original bacterial cell used to produce
`the plasmid, the DNA is collapsed by an energy-dependent en-
`zymatic reaction in which the strands are twisted into a more
`condensed, highly tensioned state. The lipids in the complex
`are thought to be in a bilayer form with the apolar lipid tails
`on the inside and the cationic head groups on the outer sur-
`face. The question is what happens when the DNA and CL are
`combined. Regular and irregular particle-like structures are
`formed with a size range of a few tens to a few hundreds of
`nanometers, but the resultant structures display significant size
`(and, presumably, compositional) heterogeneity. Whether such
`mixtures can be analyzed, even in principle, by any type of
`rigorous procedure is certainly a reasonable question. We will
`argue that they can be, although ambiguities in interpretation
`are significant.
`The actual composition of lipoplex formulations is more
`difficult to define than expected. Although only two macro-
`molecular components may be present (e.g., DNA and CL),
`they can be present either as pure species or as complexes of
`unknown composition. Furthermore, both molecules are sub-
`
`7 2 4 2 A N A LY T I C A L C H E M I S T R Y / O C T O B E R 1 , 2 0 0 7
`
`FIGURE 1. A depiction of two distinct pathways from the (left) lamellar phase to the
`(right) columnar inverted hexagonal phase of CLDCs.
`(Left) Helical DNA strands (blue ribbons) between layers of CLs (green head groups with
`yellow aliphatic chains) and neutrally charged helper lipids (white head groups with yellow
`aliphatic chains). (Center top) Along pathway I, the natural curvature C0 of the CL monolayer
`is driven negative by the addition of the helper lipid DOPE. The CL DOTAP is cylindrical,
`whereas DOPE is conical, thus leading to the negative curvature. (Center bottom) Along
`pathway II, the transition is induced by adding helper lipids consisting of mixtures of DOPC
`and the cosurfactant hexanol, which reduces the membrane bending rigidity (cid:75). (Adapted
`with permission from Ref. 17.)
`
`DNA can be hydrolyzed, and the nucleotide composition can be
`determined by LC and MS methods. A number of techniques
`exist to analyze specific degradation products as well (14).
`Perhaps surprisingly, however, the composition of CLDCs
`is not usually determined at all—rather, the composition is
`simply assumed to be defined by the identity and amounts of
`the components originally introduced. In the future, however,
`as gene therapy agents approach pharmaceutical reality, more
`rigorous criteria likely will be required.
`
`Secondary, tertiary, and quaternary structure
`Microscopy. Under certain conditions, polymers almost always
`manifest local interactions within their chains that lead to
`highly organized structure. The best known examples of this
`in biochemical systems are the (cid:65)-helices and sheets of proteins,
`the various forms of the DNA double helix, and the bilayer
`structure of amphipathic lipids. The latter two structures are
`the most relevant. The structures of DNA and CL bilayers
`have been observed both separately and in complex by X-ray
`
`

`

`diffraction; small-angle X-ray scattering; and transmission
`electron, atomic force (AFM), and scanning tunneling (STM)
`microscopies (15–20).
`From such work, a fairly uniform picture has emerged in
`which aligned strands of the helical DNA are sandwiched
`between alternating monolayers of CLs (Figure 1, left; 16).
`When neutrally charged helper lipids (so called because they
`enhance transfection efficiency) are included, a remarkable
`transformation takes place and an inverted hex-
`agonal phase of the CL part of the complex is
`formed (Figure 1, right; 17). In this state,
`the previously buried apolar side chains of
`the lipids now extend outward from the
`DNA chains, forming a uniform apolar
`coat around each strand of DNA. Al-
`though this second form of the complex
`has been accepted to transfect more
`efficiently, research has shown that fac-
`tors other than structure may account for
`differences in efficiency of gene delivery. In
`one case, increasing the charge density was
`shown to increase the transfection efficiency of
`lamellar complexes to levels observed with the in-
`verted hexagonal complex (21). A separate study showed that
`an optimum charge density may exist that balances the struc-
`tural stability of the complex with the release of the DNA (22).
`Unfortunately, the methods used to achieve these insights into
`CLDC structure are not applicable to complexes in pharma-
`ceutical formulations. They do, however, clearly establish the
`helical nature of the DNA and the local structure of the CLs.
`A quite different problem occurs when imaging methods
`are used to probe the structure of lipoplexes. Whether negative
`staining combined with electron microscopy or AFM or STM
`methods are used, one almost always sees images containing
`a collection of amorphous entities described as anything from
`nebulous globs to structures that resemble spaghetti and
`meatballs (19). At present, the utility of such data for rigorous
`analysis remains to be established. Although the helical nature
`of the DNA can sometimes be seen, typically little further
`detail is apparent.
`Dynamic light scattering (DLS). In addition to direct im-
`aging methods, the size and shape of lipoplexes can be charac-
`terized by DLS. In this method, the fluctuations in the inten-
`sity of scattered light due to the Brownian motion of particles
`are analyzed by autocorrelation methods to yield a diffusion
`constant and ultimately a size in the form of a hydrodynamic
`diameter. Although the polydispersity of the complexes formed
`by CLs and plasmid DNA makes a rigorous analysis of the
`resulting data somewhat difficult, a mean diameter can still be
`obtained or the data deconvoluted in such a way as to obtain
`distinct populations of particles as a function of approximate
`size ranges.
`When DLS is used in the presence of an electromagnetic
`field, an estimate can be made of the charge on the particle or
`the (cid:90) potential (the voltage at the surface of shear). The inten-
`sity of the scattered light as a function of the angle of detection
`
`also can be used to directly measure the molecular weight of
`the particle as well as its radius of gyration (which is based on
`the distribution of mass of the particle rather than its hydro-
`dynamic behavior). The ratio of the radius of gyration to the
`experimentally determined or calculated hydrodynamic radius
`provides a measure of the asymmetry of the particle.
`DLS studies have identified CL–DNA particles in the size
`range 50–300 nm and confirmed their intrinsic heterogene-
`ity (23–25). Their (cid:90) potentials range from the highly
`negative (an excess of DNA) to the highly posi-
`tive (an excess of CL). One of the few cor-
`relations between the physical properties of
`particles and their ability to transfect cells
`is based on DLS measurements where
`it has been found that smaller (90–150
`nm), positively charged lipoplexes gen-
`erally produce good gene expression.
`This probably reflects the mechanism by
`which such particles enter cells; initially,
`the mechanism appears to involve elec-
`trostatic interactions with highly negatively
`charged cell-surface proteoglycans (26, 27).
`Analytical ultracentrifugation. This method
`can be used in one of two modes. In equilibrium sedimenta-
`tion, material is sedimented at such a speed that equilibrium
`is reached in the centrifuge tube. The resultant distribution
`of mass of the solute in the tube is then optically monitored.
`This distribution can be analyzed and converted to an absolute
`molecular weight, which can in turn be used to calculate a size
`on the basis of assumptions about the density and shape of the
`particles. Densities (partial specific volumes) can be measured
`experimentally with oscillating U-tubes or pycnometers. In
`the second approach, the rate at which the particles sediment
`is measured (sedimentation velocity); this rate permits an es-
`timate of hydrodynamic size to be made. Unfortunately, the
`usual heterogeneity of lipoplexes has made it difficult to apply
`either method.
`A simple but less informative method is based on the sedi-
`mentation of lipoplexes in preformed density gradients of inert
`solutes, such as sucrose or dextrose (28, 29). This method has
`been used to provide empirical measures of particle behavior
`that are not related simply to their physical properties. As the
`homogeneity of pharmaceutical preparations of lipoplexes
`increases, both sedimentation velocity and equilibrium experi-
`ments have the potential to play important roles as analytical
`tools.
`Gel electrophoresis. Although sufficiently porous size-exclu-
`sion matrices exist, this method has yet to be widely applied to
`the structural characterization of lipoplexes. The gel retarda-
`tion assay is occasionally used to determine the positive-to-
`negative charge ratios that minimize the amount of unbound
`DNA (30). Because of the relative charge and size differences
`between lipoplexes and plasmid DNA, electrophoresis of the
`two produces widely separated bands. The gel retardation assay
`shows that with increasing charge ratios the amount of un-
`bound DNA decreases and the quantity of lipoplexes retarded
`
`O C T O B E R 1 , 2 0 0 7 / A N A LY T I C A L C H E M I S T R Y 7 2 4 3
`
`

`

`very concentrated and even dried materials can be examined
`easily (37, 38). In such studies, peak positions corresponding
`to vibrational modes of a wide variety of molecular groups,
`including vibrations of the DNA bases and phosphate groups,
`as well as peak positions corresponding to lipid methylene and
`carbonyl stretching bands are measured as a function of CL-
`to-DNA ratios.
`Titration of one component into the other clearly reveals
`distinct peak positions for various stages of complex forma-
`tion—this supports the use of this method for structural char-
`acterization and verification. Figure 3 presents two examples in
`which the CLs DOTAP and DDAB were titrated into a solu-
`tion of plasmid DNA. During these titrations, the positions of
`peaks corresponding to various vibrational modes and stretch-
`ing bands were monitored. The data show shifts in the peak
`positions occurring over a range of CL-to-DNA weight ratios;
`these correspond to distinct structural states in the complexes.
`Similar, but preliminary, studies have been undertaken with
`Raman spectroscopy (32), but that approach is limited because
`of the higher concentrations needed to obtain spectra.
`Fluorescence spectroscopy. This technique has been used to
`examine various aspects of lipoplex structure. The most com-
`monly used method is the simple displacement of fluorophores
`bound to plasmid DNA. Typically, dyes are either intercalated
`between the bases of the DNA or bound within the minor
`groove. The normally solvent-quenched quantum yield of fluo-
`rescent dyes is dramatically enhanced when they are bound to
`DNA. When a polycation such as a CL micelle is added to such
`labeled DNA, the fluorescent dye often is displaced through
`competition with the former CL. This displacement of the
`dye leads to a reduction in fluorescence quantum yield that is
`easily measured. If a combination of dyes is used along with
`titration experiments, lipoplex-induced structural (especially
`topological) changes in the DNA can be detected, and changes
`in the solvent accessibility of the minor groove are seen (39).
`Thus, simple measurements of dye fluorescence intensity and
`wavelength emission maximum can be used to obtain at least
`comparative structural information. The major disadvantage
`of this approach is that, unlike CD and FTIR studies, a label
`is needed. Furthermore, the label is usually added to the DNA
`before complex formation occurs. Therefore, this method can-
`not be used to characterize an intact lipoplex directly.
`Another fluorescence-based method that suffers from some
`of the same problems but is of significantly higher resolution
`is fluorescence resonance energy transfer (FRET). When two
`fluorescent groups have the property that the emission spec-
`trum of the donor overlaps the absorption spectrum of the ac-
`ceptor, it is possible to excite the donor but see emission from
`the acceptor (or quenching of the donor; 40). By measuring
`the efficiency of such events, one can estimate the distance
`between the donor(s) and acceptor(s).
`FRET is a versatile technique that is increasingly seen in
`a wide variety of applications (41). In the case of lipoplexes,
`it has been possible to label the DNA by either intercalation
`of a dye between the bases or binding of a dye to the minor
`groove (donors). Then, a different dye can be placed within
`
`near the top of the gel increases. Complete retardation of DNA
`often does not occur until an excess of positive lipid charge to
`negative DNA charge exists; this probably indicates that not
`all of the positive charge associated with the lipid is involved
`in the formation of the complex. Farhood et al. attributed this
`observation to steric hindrance between the two species due to
`the bulky nature of the liposome and superhelical DNA (31).
`
`DNA
`0.25
`0.5
`1.0
`1.5
`
`DNA
`0.25
`0.5
`1.0
`
`1.5
`
`10,000
`
`5000
`
`0
`
`–5000
`
`–10,000
`
`(cid:2)(cid:1)(degrees L mol –1 cm –1)
`
`200
`
`220
`
`240
`
`300
`280
`260
`Wavelength (nm)
`
`320
`
`340
`
`FIGURE 2. Representative CD spectra of DOTAP–DNA complexes.
`The charge ratios are indicated. Open symbols are CLDCs below charge
`neutrality, and the closed symbol is a CLDC above charge neutrality.
`(Adapted with permission from Ref. 22.)
`
`Circular dichroism (CD). This straightforward technique
`focuses primarily on the helical structure of DNA by measuring
`the difference in absorption of left- and right-handed circularly
`polarized light. Because of the regular helical array of the
`DNA bases, very strong optical activity is present within DNA
`molecules. For the most common type of DNA, known as B-
`form, a very characteristic spectrum displays a strong positive
`peak near 275 nm and a negative signal of similar intensity at
`~245 nm. When CLs are added to DNA, a red shifting of both
`peaks and a fairly dramatic reduction in intensity of the 275 nm
`feature are seen (Figure 2). These large spectral changes were
`originally thought to be due to a change from B-form DNA (10
`bp/turn) to C-form (9 bp/turn). Later studies, however, based
`on IR and Raman measurements and molecular dynamics
`simulations, strongly suggest that the changes seen are due to
`small alterations in the interactions between the bases while the
`DNA essentially remains in the canonical B-form (32). What is
`important from an analytical point of view is that the CD spec-
`tra of the complexes are very sensitive to their structure and that
`therefore CD can monitor the state of the DNA in lipoplexes
`and polyplexes (cationic polymer-based complexes; 33–35).
`FTIR. This method can characterize the structure of both
`the DNA and CL components of complexes (36). A disadvan-
`tage of this technique is that 10–20-fold higher concentra-
`tions are required for solution studies; an advantage is that
`
`7 2 4 4 A N A LY T I C A L C H E M I S T R Y / O C T O B E R 1 , 2 0 0 7
`
`

`

`(a)
`
`(b)
`
`(c)
`
`2926
`2925
`2924
`2923
`2855
`2854
`2853
`2852
`
`1746
`1742
`1738
`1734
`
`(d)
`
`(e)
`
`(f)
`
`1720
`1718
`1716
`1714
`
`1492
`
`1490
`
`1488
`
`1227
`1225
`1223
`1221
`
`0
`
`(g)
`
`(h)
`
`2923
`2921
`2919
`2917
`
`2853
`2852
`2851
`2850
`
`(i)
`
`1717.5
`1716.5
`1715.5
`1714.5
`
`(j)
`
`(k)
`
`1493
`1491
`1489
`1487
`
`1227
`1225
`1223
`1221
`
`2
`1
`Weight ratio (CL/DNA)
`
`3
`
`0
`
`3
`2
`1
`Weight ratio (CL/DNA)
`
`4
`
`Wave number (cm–1)
`
`the bilayer of the CLs (acceptors); this makes
`it possible to estimate the mean distance be-
`tween various sites within the complexes (23,
`42). Although they have only been explored
`to a limited extent, versions of this approach
`may turn out to be quite useful. For example,
`FRET measurements with respect to time
`can be used to obtain the kinetics of complex
`formation even at very early times, permitting
`one to see critical structural rearrangements
`as complexes are created (43, 44).
`
`Stability of complexes
`If lipoplexes are to be used as pharmaceutical
`agents, methods to characterize their stability
`in real time (storage) and accelerated modes
`will be necessary. The idea is that by increas-
`ing temperature, changing pH, agitating the
`complexes, or applying other stresses, one
`can increase degradation to the point that it
`can be detected at short time intervals and
`the drug’s environment adjusted accordingly.
`In addition to the methods described below,
`these changes can be determined by the
`methods described earlier (36, 45–48).
`Gel electrophoresis. Agarose gel electro-
`phoresis combined with intercalating dyes is
`a common method to visualize the size and
`conformation of plasmid and linear DNA.
`Plasmid DNA commonly exists as three iso-
`forms (linear, open circular, and supercoiled)
`that migrate through the gel at different rates
`based on their shapes. Because plasmid DNA
`undergoing oxidative degradation often
`shows a transition from predominantly su-
`percoiled to open circular and eventually to
`linear forms because of cleavage of the phos-
`phodiester backbone, monitoring the change in the relative
`amounts of the isoforms can serve as a simple stability assay.
`The stability of intact complexes has also been characterized
`with the gel retardation assay described earlier (49). Complexes
`that have undergone increasing levels of degradation produce
`bands corresponding to more unbound DNA.
`A related method can be used to characterize the capacity
`of the CL to protect the genetic material from degradative
`enzymes that are likely to be encountered inside and outside
`the cells (50). Here, lipoplexes are exposed to DNase I, which
`cleaves unbound and/or unprotected DNA into linear frag-
`ments. A detergent is then used to separate the DNA from
`the CL, and the components are detected by agarose gel
`electrophoresis.
`Chemical probes. DNA can organize into a variety of
`complex secondary structures, such as left-handed DNA, cru-
`ciforms, and other multistranded structures (51). A number
`of techniques that use chemical probes have been developed
`to recognize and detect minor changes within the secondary
`
`FIGURE 3. The effect of (a–f) DOTAP:DNA and (g–k) DDAB:DNA weight ratios on DNA
`and CL vibrational modes.
`The peak positions were plotted versus the weight ratio of CL to DNA with 1 mg/mL DNA
`in each sample. Stretching bands: (a, g) lipid asymmetric methylene; (b, h) lipid symmetric
`methylene; (c) lipid carbonyl; (d, i) guanine/thymidine carbonyl; (e, j) guanine/cytosine, in-
`plane; (f, k) DNA asymmetric phosphate. (Adapted with permission from Ref. 37.)
`
`structure (52, 53). One such probe, KMnO4, selectively oxi-
`dizes unpaired thymine residues, thus making the DNA sus-
`ceptible to piperidine cleavage of the modified phosphodiester
`backbone. This is resolved with polyacrylamide gel electro-
`phoresis. In contrast to a bulky enzymatic probe, the relatively
`small size of KMnO4 allows detection of changes as minor as
`strand separation at the base pair level and can therefore be
`used to detect DNA melting within a lipoplex. One such study
`found that upon initial interaction between DNA and CLs, the
`DNA undergoes a partial unwinding (54).
`Isothermal titration (ITC) and differential scanning
`(DSC) calorimetries. One indirect method that has been ex-
`tensively used to characterize the stability of complexes is ITC
`(55–57). In this approach, a concentrated solution of either
`the DNA or the CL is added in small increments to the other,
`and the small amount of heat released or absorbed is measured
`by compensating for the small temperature changes produced
`by the binding interaction. The results are analyzed by fitting
`the resultant data to various binding models, and in principle,
`
`O C T O B E R 1 , 2 0 0 7 / A N A LY T I C A L C H E M I S T R Y 7 2 4 5
`
`

`

`120
`
`for short pieces of DNA, the more extensive interactions
`that occur within a plasmid molecule (many thousands
`of DNA base pairs in length) produce a very high Tm
`(>90 °C).
`The results of DSC experiments with lipoplexes pro-
`duce the transitions expected for the CL and DNA
`components (58). In the case of complexes that contain
`CLs that undergo phase transitions within a measurable
`range, a sharp transition (Figures 4d and 4g) is typi-
`cally seen. When the bilayer is stabilized by the DNA,
`this transition is usually shifted to a higher temperature
`(Figures 4b to 4c and 4e to 4f).
`This phase transition also can be seen directly with
`IR spectroscopy by analyzing the methylene stretch-
`ing vibrations or through the use of fluorescent probes
`which insert into the bilayer. A broader transition is
`seen at >100 °C that reflects the melting of the plasmid
`(Figures 4a to 4c, 4e to 4f, and 4h to 4i). It is possible
`to go above 100 °C, because the experiments can be
`conducted under pressure, which will raise the boiling
`point of water. A complex third transition is also seen at
`60–70 °C, which reflects the presence of contaminat-
`ing linear and open circular forms of DNA (Figure 4a).
`Both high-sensitivity ITC and DSC are now available in auto-
`mated modes; this makes them applicable to high-throughput
`lipoplex analyses.
`UV absorption. Because the absorption of UV light is
`sensitive to changes in base pair interactions, the absorption
`spectrum of DNA strongly reflects the degree of secondary
`structure formation. Recent technical advances in the use of
`diode array spectrophotometers combined with computer-
`aided data analysis now permit derivative spectra of ±0.01 nm
`resolution to be easily obtained, thereby allowing signals from
`the different types of bases to be resolved. Because of base pair
`interactions that occur within the ordered secondary structure
`of double-stranded DNA, UV absorption is less than that
`predicted solely on the basis of nucleotide concentrations (45).
`As DNA undergoes a loss of secondary structure (melting),
`UV absorption increases. Monitoring absorption as a function
`of temperature is especially useful for the generation of DNA
`melting curves. One can quickly and easily generate melting
`curves and measure the Tm of different lipoplexes over a range
`of charge ratios with a variety of automated instrumenta-
`tion. For example, Zhang and colleagues showed that the CL
`DDAB increased the Tm of double-stranded calf thymus DNA
`by ~8 °C (59).
`Empirical phase diagrams (EPDs). The analysis and char-
`acterization of lipoplexes are difficult because of their intrinsic
`complexity and heterogeneity. In the case of better-defined
`pharmaceuticals, facile methods such as the various types of
`HPLC are quite useful. Methods such as MS and NMR can
`provide very detailed structural pictures because of their high
`information content. Their current applicability to lipoplexes
`is, however, low. In contrast, lipoplexes require analysis by a
`wide variety of methods that probe various aspects of their
`structure and stability. Multiple measurements have the po-
`
`i
`
`h
`
`f e
`
`c b
`
`a
`
`g
`
`d
`
`60,000
`
`50,000
`
`40,000
`
`30,000
`
`20,000
`
`10,000
`
`Molar heat capacity (kcal/K mol)
`
`0
`40
`
`50
`
`60
`
`80
`70
`90
`Temperature (°C)
`
`100
`
`110
`
`FIGURE 4. DSC thermograms of CLDCs and cationic liposomes alone.
`(a) DNA; (b, c) DDAB–DNA complexes at positive-to-negative charge ratios of
`(b) 0.75:1 and (c) 1.0:1; (d) DDAB liposomes alone; (e, f) distearoyltrimethylam-
`monium propane (DSTAP)–DNA complexes at (±) ratios of (e) 0.75:1 and (f) 1.0:1;
`(g) DSTAP liposomes alone; (h, i) DOTAP–DNA complexes at (±) ratios of (h) 0.75:1
`and (i) 1.0:1. (Adapted with permission from Ref. 58.)
`
`the equilibrium constant, stoichiometry (molecular ratio of the
`two components in the complex at saturation), enthalpy, and
`entropy of binding are determined. Unfortunately, the interac-
`tions between a plasmid DNA molecule and a CL particle are
`complex; this makes the determination of thermodynamic pa-
`rameters difficult, if not impossible. Irreversibility of binding,
`aggregation, and structural changes imply a lack of equilibrium
`and the occurrence of multiple events—situations that pro-
`hibit an unambiguous analysis of ITC data. Furthermore, this
`method is not applicable to preformed complexes, the desired
`targets of most stability studies.
`Another form of calorimetry, DSC, is of direct utility for
`stability analyses. The lipoplexes are heated simultaneously
`with a reference sample (buffer), and sufficient thermal energy
`is supplied to maintain the solutions at the same temperatures.
`If a change in structure occurs in the target material, the com-
`pensating energy required to maintain the same temperature
`in the two samples can be converted to an energy associated
`with the structural alteration. In the case where the transition
`is reversible, the enthalpy of the structural alteration can be
`obtained. Although this condition is usually not the case, it is
`still possible to measure a “melting temperature” Tm, which is
`related to the thermal stability of the complex.
`Two types of thermal transitions can occur within lipoplexes.
`The first corresponds to a transition from the gel phase to the
`liquid crystal phase; during this process, the ordered alkyl side
`chains of the CLs are converted at elevated temperature to a
`more fluid state. The phenomenon may or may not be displayed
`by a particular CL bilayer. The second type of transition is pro-
`duced by the melting of the two chains that make up the DNA
`double helix. These two strands are held together primarily by
`hydrogen bonds between the bases as well as by interactions be-
`tween their planar surfaces. Although this disruption of DNA
`structure is expected to occur at low temperatures (25–60 °C)
`
`7 2 4 6 A N A LY T I C A L C H E M I S T R Y / O C T O B E R 1 , 2 0 0 7
`
`

`

`4
`
`5
`
`6
`pH
`
`4
`
`5
`
`6
`pH
`
`150
`125
`100
`75
`50
`25
`10
`
`150
`125
`100
`75
`50
`25
`10
`
`Ionic strength (mM)(
`
`a)
`
`Ionic strength (mM)(
`
`c)
`
`FIGURE 5. Ionic-strength–pH EPDs of various nonviral gene delivery complexes generated from DLS,
`CD, and fl uorescence studies.
`(a, c) DOTAP lipoplex and (b, d) DOTAP–DOPE lipoplex at positive-to-negative charge ratios of (a, b) 0.5 and
`(c, d) 4. Regions of similar color represent single phases, whereas the change of color defi nes the conditions
`under which the structure of the gene delivery complex alters. (Adapted with permission from Ref. 25.)
`
`150
`125
`100
`75
`50
`25
`10
`
`Ionic strength (mM)(
`
`b)
`
`7
`
`8
`
`4
`
`5
`
`6
`pH
`
`7
`
`8
`
`150
`125
`100
`75
`50
`25
`10
`
`Ionic strength (mM)(
`
`d)
`
`7
`
`8
`
`4
`
`5
`
`7
`
`8
`
`6
`pH
`
`at positive-to-negative charge ratios of 0.5 and 4.0 (Figure 5).
`The resultant 5D, colored EPDs display three to five vari-
`ably resolved regions that correspond to lipoplexes displaying
`structures with varying degrees of size, DNA collapse, and sec-
`ondary structure. Interpretation of the colored regions within
`the EPD, however, requires analysis of the original data (25).
`The light blue-green regions observed at pH 7 for DOTAP
`lipoplexes (Figure 5c) and at pH 7 and 8 for DOTAP–DOPE
`lipoplexes (Figure 5d) at high charge ratios ref