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www.chromatographyonline.comApril 2014 Volume 32 Issue s4

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Recent Developments in

LC ColumnTechnology

Recent Developments in

LC ColumnTechnology

6 RECENT DEVELOPMENTS IN LC COLUMN TECHNOLOGY APRIL 2014

Articles

HPLC Column Technology 2014: State of the Art . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8Ronald E. Majors

A brief introduction to the articles presented in this supplement

Impact of Particle Size Distribution on HPLC Column Performance . . . . . . . . . . . . . . . . . . . . . . . . . 12Richard A. Henry

Controlling particle size distribution is examined as a possible route to further improve the performance of particle-based columns.

Recent Progress in Chiral Stationary Phase Development and Current Chiral Applications . . . . . . . . 20Timothy J. Ward and Karen D. Ward

A review of chiral separations, which remain a decided area of interest, particularly in the pharmaceutical and agrochemical fields

The Role of Chromatography in the Characterization and Analysis of Protein Therapeutic Drugs. . . . 24C. David Carr

Chromatography has taken a prominent place in the characterization and analysis of protein

therapeutic drugs and today it plays a critical role in the biotechnology laboratory.

Size-Exclusion Chromatography of Protein Aggregation in Biopharmaceutical Development and Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30Linda Lloyd

Size-exclusion chromatography is an important technique in biopharmaceutical characterization.

This article discusses its use for soluble aggregation analysis and quantitation.

Impact of New Columns on Drug Development. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36Joseph L. Glajch

An examination of the development of new types of columns based on different particle

types, sizes, and other physical characteristics and how they can improve the speed and

efficiency of HPLC used to support more expansive and complicated analyses

Liquid Chromatography Methods for the Separation of Short RNA Oligonucleotides. . . . . . . . . . . . . 42Mirlinda Biba, Bing Mao, Christopher J. Welch, and Joe P. Foley

A review of LC methods and strategies for the chromatographic separation of short RNA oligonucleotides

Apr i l 2014

Volume 32 Number s4

Cover images courtesy of LAGUNA DESIGN/Shunyu Fan/Tuomas Marttila/PASIEKA/Getty Images; Dan Ward


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N O W Y O U C A N S TA R T T H I N K I N G O F

N O T R E S T R A I N T S .

I N T E R M S O F


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Ronald E. MajorsLCGC columnist and analytical consul-tant, West Chester, Pennsylvania. Direct correspondence to: [email protected]

HPLC Column Technology 2014: State of the Art

In this special supplement to LCGC North America, you will find the

latest updates on high performance liquid chromatography (HPLC)

and ultrahigh-pressure liquid chromatography (UHPLC) column

technology. This year’s supplement has an emphasis on columns

for both small molecules and macromolecules, particularly those

developed for the biopharmaceutical and biotechnology industry.

Since the last supplement on liquid chromatography (LC) column tech-nology in 2012, the continued inter-

est in high-throughput separations and pro-ductivity increases has been a strong driving force in column development. The rapid adaptation of superficially porous particle (SPP) columns, also referred to as core–shell or fused-core particle columns, has been phe-nomenal. In my recent Pittcon 2014 article (1), in terms of new product introductions, it seems the entire columns world has turned to SPP as the favored type of high perfor-mance liquid chromatography (HPLC) or ultrahigh-pressure liquid chromatography (UHPLC) column, with SPP introductions exceeding sub-2-µm columns by a 10:1 mar-gin. It is no wonder — the most popular SPP columns of 2.6–2.7 µm particle size provide the same efficiency as the sub-2-µm columns (1.7–1.9 µm) yet present only half of the back pressure. In addition, the introduction of larger SPP particles up to 5 µm, including larger-pore versions for protein and peptide separations, have been noted as well as sub-2-µm SPP versions, which offer phenomenal efficiencies at the expense of higher back pressure. Because many users have upgraded to UHPLC instruments with pressure capa-bility up to 19,000 psi, the increased pres-sure requirements did not constitute a large burden. A bigger problem facing instrument companies is to provide low-dispersion flow systems that can bring out the potential ultrahigh efficiency of these small-particle SPP columns.

This issue is devoted to columns that are particularly suited to the pharmaceuti-cal industry. In particular, the overwhelm-ing interest in biological-based drugs has spawned further interest in improvements of columns that can handle large biomolecules such as proteins and nucleic acid species. In addition, a new industry of biosimilars is popping up that presents challenges to the regulatory bodies in providing drug safety and efficacy information where chromatog-raphy is a key analytical tool.

In terms of the rapid changes occurring in the biopharma world, all one has to do is to look at the total sales of pharmaceuticals in the last few years. Figure 1a shows a chart of the top 200 drugs in terms of revenue in 2006, only a short 8 years ago when bio-pharmaceuticals were already in progress. (I don’t expect you to read the details of these charts but to look at the number of those boxes highlighted in green; for high resolu-tion charts go to reference 3.) In that year, out of a total of 200 top-selling drugs only 19 were drugs of biological origin. A similar chart for 2012 is depicted in Figure 1b. In this chart there are a total of 50 drugs that fit the biological category, more than 2.5 times the number compared to only 8 years ago. It is expected that this trend will continue.

To update LCGC readers in columns for new drug entities, I have assembled an excel-lent selection of separation science experts to bring us up-to-date in specific areas of stationary phase technology. First, Dick Henry, well known to many of us in the

LAGUNA DESIGN/SHUNYU FAN/TUOMAS MARTTILA/PASIEKA/GETTY IMAGES; DAN WARD


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separation science field, discusses the role of particle size distribution in chromatography efficiency and pressure. There has been a lot of discussion recently about whether or not the narrow particle distribution of SPPs is mainly responsible for the efficiency gains for these particles, but Dick will show you that even totally porous particles can gain from having a narrow size distribution. The team of Tim and Karen Ward of Milsaps College, experts in chiral column technol-ogy, update us on their 2012 article (4) covering new columns for this important area of pharmaceutical analysis. It is well known that enantiomeric purity can con-trol the effectiveness of administered drugs where one racemate is beneficial while the

other racemate can cause side effects or even worse.

Next, Dave Carr of Bioanalytical Tech-nologies, who has been teaching about char-acterization and analysis of protein therapeu-tic drugs for years, discusses the role of new columns in tackling difficult separations of polypeptides, peptides, and proteins by a variety of modes including reversed-phase chromatography, ion-exchange chromatog-raphy, and size exclusion-chromatography (SEC). In the following article written by Linda Lloyd, a biocolumns product manager at Agilent Technologies, the separation tech-nique of SEC is examined more deeply with an emphasis on protein aggregation. She looks at how SEC can provide analytical data

in upstream and downstream production as well as play a major role in the formulation studies and quality control of drug sub-stances and drug products such as monoclo-nal antibodies. Joe Glajch of Momenta Phar-maceuticals brings an industrial perspective on the impact of new columns on drug development, particularly with his involve-ment in the biosimilars. He discusses the use of monoliths, sub-2-µm porous particle, and SPP columns in his laboratory with exam-ples of rapid N-glycan, tryptic peptide, and amino acid, and heavy and light chain vari-ants of an IgG monoclonal antibody analysis.

Last but not least, Mirlinda Biba and coworkers from Merck tackle the relatively new area of oligonucleotide-based pharma-ceuticals. Biba is in the process of getting her PhD from Drexel University and is well versed in the needs of the high purity require-ments for the development of oligos destined to become potential therapies for the treat-ment of disease from her work at Merck.

I hope that you enjoy this special supple-ment and find something of interest and value that may help you in solving your liquid-phase separation problems.

References

(1) R.E. Majors, LCGC North Am. 32(4), 242–

255 (2014).

(2) J.T. Njardarson et al., J. Chem. Ed. 87, 1348

(2010).

(3) Charts courtesy of the Njardarson Group, Cor-

nell University, M. Brichacek, N. McGrath,

E. Rogers, J.Morton, L.Batory, R. Bauer, J.A.

Wurst, and J.T. Njardarson. To obtain high

resolution versions of these charts, please refer

to the following URL: http://cbc.arizona.edu/

njardarson/group/top-pharmaceuticals-poster.

(4) T.J. Ward and K.D. Ward, LCGC North Am.

30(s4), 43–45 (2012). �

Figure 1: Top 200 pharmaceutical products by United States retail sales (a) in 2006 and (b) in 2012. Adapted from references 2 and 3.

Ronald E. Majorsis the editor of “Column Watch” and “Sample Prep Perspectives.” He is also an analytical consultant and a member of LCGC’s editorial advisory board. Direct correspondence about this column to [email protected]

For more information on Ron Majors or to see his past columns, please visit

www.chromatographyonline.com/majors


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Richard A. HenryConsultant, State College, Pennsylvania.Direct correspondence to: [email protected]

Impact of Particle Size Distribution on HPLC Column Performance

In this article, controlling the particle size distribution (PSD)

variable has been examined as a possible route to further

improvement in performance of particle-based columns.

Si l ica part icles prepared in tubular column formats domi-nate high performance liquid

chromatography (HPLC), and the technique would not be as important without them. Over the years, silica particles have been optimized in shape, purity, and size. They have steadily become smaller to improve efficiency, which allows for more speed, sensi-tivity, and resolution at the cost of higher pressure. Knox (1) and others published early papers showing accu-rate insight into particle and pressure requirements for improving HPLC speed and performance.

When discussing column perfor-mance, it is always helpful to start with the van Deemter relationship, which describes three additive pro-cesses that inf luence bandspreading within the column.

H = A + B/µ + Cµ [1]

H stands for height equivalent to one theoretical plate (HETP) and is calcu-lated from a measurement of column efficiency, N, and the relationship, H = L/N, where L is column bed length. The µ term is the linear velocity of the mobile phase. A plot of H versus µ is commonly called a van Deemter curve and represents how efficiency behaves

as a function of µ (proportional to f low rate). The lower the H value, the greater eff iciency a column displays. In equation 1, the A term represents contributions from f low and diffusion processes within the mobile phase f lowing around the particles (names include eddy diffusion, eddy disper-sion, f low inequality, and multipath term), the B term is total axial (lon-gitudinal) diffusion within mobile and stationary phases, and the C term represents speed of mass transfer between mobile phase and stationary phase that lies mainly within particle pores. If a narrower particle distribu-tion can create more uniform beds, it should show up as a smaller A term, which is not very sensitive to f low velocity compared to the other two terms. The A term dominates in the middle of the velocity curve and con-trols the lowest value of plate height at which efficiency of a column is high-est, while the B term dominates at low velocity and the C term dominates at high velocity. A complete discussion of factors affecting band dispersion is beyond the scope of this article and for an in-depth discussion of van Deemter relationships, refer to Neue (2) and other excellent HPLC books.

Typical van Deemter curves are shown in Figure 1 for several sizes of


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porous C18 silica to demonstrate the two-fold advantage of selecting smaller particles (3). Not only can smaller H (larger N ) be achieved by using smaller particles, this better column performance can be sustained even at higher velocities (smaller slope for the C term). Note that Figure 1 uses interstitial linear velocity rather than

classical linear velocity, which is more easily obtained from the relationship, µ = L/t0, where L is column bed length and t0 is the time for an unretained peak. Neue (2) and other texts provide definitions of different linear velocity terms and other HPLC column fun-damentals. Plots of H (or h, called the reduced plate height, H/dp where dp

is the average particle diameter in the same units as H ) against classical lin-ear velocity are usually adequate and will be used in this article. Reduced plate height allows performance to be compared when columns use different particle sizes. Until modern core-type particles were introduced in 2006, the lowest hmin values observed for silica particles were 2–3.

A lthough Figure 1 creates the impression that nearly unlimited sep-aration speed might be possible with sub-2-µm particle columns, column resistance and instrument pressure rating quickly become limiting factors. Measurement of H-µ (or h-µ) curves with simple, ideal solutes is very use-ful in column research; however, real samples contain solutes that may vary greatly in curve shape. When high speed methods are being developed, kinetic performance with H-µ plots should be compared for both ideal and target solutes during column screen-ing. For example, the best performing column with toluene or naphthalene may not yield equally good perfor-mance for drug metabolites. Plots of column efficiency, N, and resolution, R, against linear velocity can also be very valuable during the development of a high-speed method. Column sta-bility toward high f low and pressure conditions should also be confirmed. Figure 2 illustrates how well modern HPLC columns can operate at very high linear velocity (circa 20 mm/s) under gradient conditions (4). Gradi-ents are often used in high-throughput assays to clean the column between sample injections. The 20 mm × 2.1 mm C18 column prepared with 5-µm core-type silica particles required a starting system pressure of 172 bar (78 bar from the column and 94 from the ultrahigh-pressure liquid chroma-tography [UHPLC] instrument).

New insight about van Deemter relationships was provided by Knox (5,6), who suggested that the A term is much more important than previ-ously thought and should receive more attention as a path to improved HPLC column performance. According to Knox, a lower A term will come from better column preparation techniques and more uniform bed structures. In

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Figure 1: A van Deemter plot for small porous particles. Columns: 50 mm × 4.6 mm Zor-bax Eclipse XDB-C18 (1.8-µm column was 30 mm in length); eluent: 85:15 acetonitrile–wa-ter; �ow rates: 0.05–5.0 mL/min; temperature: 20 °C; sample: 1.0 µL octanophenone in eluent. (Courtesy of Ron Majors and Agilent Technologies.)

Figure 2: Fast gradient assay with a 5-µm core-type column. Column: 20 mm × 2.1 mm, 5-µm dp Ascentis Express C18; mobile-phase A: water with 0.1% tri�uoroacetic acid; mo-bile-phase B: acetonitrile with 0.1% tri�uoroacetic acid; mobile phase: 11:89 A–B; �ow rate: 2 mL/min; pressure: 172 bar; temperature: 40 °C; detection: UV absorbance at 254 nm; injection volume: 0.5 µL; �ow cell: 1-µL micro. Peaks: 1 = atenolol, 2 = pindolol, 3 = propranolol, 4 = indoprofen, 5 = naproxen, 6 = coumatetralyl. (Courtesy of Advanced Materials Technology.)


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APRIL 2014 RECENT DEVELOPMENTS IN LC COLUMN TECHNOLOGY 15www.chromatographyonline.com

this article, the particle size distribu-tion (PSD) variable has been exam-ined as a possible route to higher per-formance for particle-based columns.

Experimental

HPLC column preparat ion can involve a huge number of variables, making it diff icult to draw accurate conclusions from experiments with particles unless preparation variables are optimized or at least held con-stant. Another important variable for studying high performance columns is selecting a suitable instrument that measures true bed efficiency by mini-mizing extracolumn bandspreading. This article focuses on the impact of PSD and assumes that variables such as column preparation and testing have been controlled and optimized to an acceptable degree.

Different methods for measuring and reporting particle size and dis-tribution are described by Bartle and Myers (7) and Horiba (8). Unless otherwise specif ied, this article will use the common D 90/10 method for reporting distribution. D90/10 is defined as the particle diameter (dp) at 90% of the distribution (a larger number) divided by dp at 10% of the distribution (a smaller number). Par-ticles used for preparing HPLC col-umns can range from values less than 1.1 to 1.5 or more. Figure 3 shows distribution data for six spherical porous silica samples taken from com-mercial HPLC columns. As shown in the next section, column performance with spherical particles in broader size ranges is still quite good and allows the HPLC technique to solve many important analytical problems.

Results and Discussion

Important benefits to achieving even more uniform column beds should include providing very consistent col-umn performance and greater operat-ing stability. A near-perfect bed should be resistant to settling or voiding and would presumably fail only because of contamination or loss of stationary phase (a factor of temperature and pH). While progress in silica particle devel-opment continues, parallel efforts are underway to accomplish similar per-formance and stability objectives using monolithic silica beds and fabricated pillar arrays. Results described in this article have been taken from both pub-lished and unpublished sources.

Verzele (9) investigated C18 column performance under reversed-phase con-ditions using spherical silica samples having relatively narrow distribution (D90/10 = 1.5). He blended samples to deliberately broaden the distribution and measured column performance before and after blending. Verzele did not observe significant differences in hmin between blended and unblended samples, but he did note that particles blended to create broader distribu-tion showed greater f low resistance (pressure drop) and lost performance slightly faster at higher f low veloc-ity (higher slope for C term). Imped-ance reached a maximum with a 50/50 blend of 8- and 3-µm particles that was about 50% greater than a column pre-pared with only narrow-distribution 3-µm particles. Verzele recommended that HPLC columns be prepared with D90/10 distributions of 1.5–2.0. Impor-tant conclusions that can be drawn from these early experiments on PSD is that good columns can be prepared with broader distribution samples, but at the price of higher pressure. This is an important point because pressure has become a critical factor in achiev-ing higher separation speed with sub-2-µm particle columns.

Kirkland pioneered core-type silica particles (10–13) and described the first narrow-distribution silica in 2007 (14,15), which showed extremely low hmin values of about 1.5. The particle standard deviations were very narrow at about 5% relative standard deviation (RSD) (equivalent to a D90/10 value of

16

1.14

1.141.17

1.431.511.52

Particle size (�m)

Titan sub-2.0 µm1.7-µm fully porous2.7-µm fused core3.0-µm fully porous4.7-µm fused core5.0-µm fully porous

D (90/10) shown above pro�les

14

12

10

8

6

4

2

-2

00 1 2 3 4 5 6

Figure 3: Size distribution for several silica HPLC column particles. The D90/10 value is shown above the pro�les. (Unpublished data supplied by Supelco division of Sigma-Aldrich.)

Table I: Pressure comparison for particles with different distributions: three porous particles with same average size* (courtesy of Supelco division of Sigma-Aldrich)

Pressure Profile (bar)

Instr. background 1.8 13.3 28.1 60.6 78.2

Titan C18, 1.9 µm 26.0 123.9 245.4 466.8 580.5

Column A C18, 1.9 µm 34.4 161.1 317.2 596.7 732.5

Column G C18, 1.9 µm 32.0 161.4 319.0 614.7 759.5

Velocity (mm/s) 0.43 2.17 4.35 8.52 10.68

Flow (mL/min) 0.1 0.5 1 2 2.5

* Unpublished data supplied by Supelco division of Sigma-Aldrich; 50 mm × 3.0 mm columns tested in 60% acetonitrile. (See Figure 4b for particle distribution details.)


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1.1) compared to typical values for other HPLC particles at that time of about 15–20% RSD (equivalent to D90/10 val-ues of about 1.4). Columns with greatly improved performance could be pre-pared with larger 2.7-µm particles and used with traditional 400-bar instru-ments, which was welcome news to those faced with changing to sub-2-µm particles and expensive UHPLC instru-ments. Since that breakthrough in par-ticle design, there has been considerable effort to determine why columns with core-type particles exhibit such high

performance compared to columns with porous particles. Core-type particles showed improvements (lower values) over totally porous particles in all three parts of the van Deemter equation, with the most significant advantage being in the A term, which is usually associ-ated with bed uniformity. The much narrower particle distribution might be a factor in creating such a low A term; however, Guiochon (16) proposed that other particle properties such as surface roughness are a more likely cause than narrow PSD for the highly uniform beds.

The impressive performance of core-type particles reopened the debate among academic scientists about the importance of narrow particle size dis-tribution to column efficiency. Desmet (17) studied four porous and three core-type commercial particles having about 3 µm diameter (distribution ranged from about 5% to 25% RSD) and reported strong correlations of about 0.9 between particle distribution and several differ-ent measures of column performance. Measurements that Desmet related to bed quality included the following: hmin, the A term constant, and column impedance. Based on fitted-line plots, hmin ranged from 1.5 to 2.5; the A term ranged from about 0.6 to 1.2; and sepa-ration impedance ranged from 1000 to 2500. Core-type particles with the low-est RSD had the highest performance, while the porous packing with lower RSD had higher performance within the porous group. Desmet noted that the extra performance of core-type par-ticles could also be influenced by more subtle features such as higher particle density and surface roughness. Desmet suggested that porous particles with the same narrow distribution of core-type particles should be developed and studied. Desmet (18) also examined the impact of adding 3-µm porous particles to narrow-distribution 1.9-µm porous silica and concluded that the addition of larger particles could not be expected to improve column kinetic performance because of pressure elevation; however, he reported that up to 25% of the larger particles could be added without having a major negative impact on band broad-ening and efficiency. This suggested that narrow PSD alone may not be responsible for the high performance of core-type particles. Guiochon (19) con-cluded in a different study blending 3- and 5-µm porous particles that, as long as the distribution did not exceed 40% RSD, it did not have a negative impact on band broadening and efficiency. He further proposed that intentionally adding some larger particles to smaller ones to broaden the distribution could actually improve column efficiency for small molecules, but did not discuss the negative impact that a broadened PSD might have on column permeability and kinetic performance.

Figure 4: Performance comparisons for (a) two particles with narrow distributions and one with broader distribution and (b) three porous particles with the same aver-age size and different distributions. (a) Toluene data for 50 mm × 3.0 mm columns with 60:40 acetonitrile–water mobile phase. (b) Test conditions shown in the �gure. (Unpublished data supplied by Supelco division of Sigma-Aldrich.)

12.000(a)

12.000

10.000

10.000

8.000

8.000

6.000

6.000

Mobile phase velocity (mm/s)

More monodisperse 1.9 �m and 2.7 �m

More polydisperse 1.7 �m

Titan C18 1.9 µm

Competitor B C18 1.7 µm

Ascentis Express C18 2.7 µm

D90/10

= 1.14 (both particles)

D90/10

= 1.524.000

4.000

2.000

2.000

h

0.0000.000

11.000

Columns:50 mm x 3.0 mm porous C18Test conditions: 60% acetonitrile

Column G 1.9 �m, D90/10

ca. 1.4Column A 1.9 �m, D

90/10 ca. 1.4

Titan 1.9 �m, D90/10

ca. 1.1

9.000

7.000

5.000

3.000

1.000

0.00 2.00 4.00 6.00 8.00 10.00 12.00

Mobile phase velocity (mm/s)

Column G TolueneColumn G NaphthaleneColumn G BiphenylColumn A TolueneColumn A NaphthaleneColumn A BiphenylTitan TolueneTitan NaphthaleneTitan Biphenyl

h

(b)


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Barber (20) noted that previous studies into how particle properties can impact column performance had focused on particles having the same shape. He described the blending of narrow PSD (7% RSD) porous silica spheres with twins or dimers to determine how much dimer could be present in the silica process before it negatively impacted column performance. Barber observed that up to 20% of the larger dimer particles could be added without observing a negative impact on efficiency. Contrary to results of previous experi-ments with blending spheres to broaden distribution, column permeability actually improved when dimers were blended with spheres, suggesting that the presence of some dimers was beneficial.

Henry (21) described results for the sub-2-µm porous silica shown in Figure 3 that has the same narrow distribution as core-type particles (D90/10 = 1.14). Figure 4a compares plots of reduced plate height, h, for narrow distribution porous and core-type silica particles to broader distribution porous silica. While sampling only a limited number of commercial columns, a lower overall h value observed for monodisperse, porous silica over polydisperse, porous silica supports the argument that narrow PSD can provide a performance edge (for toluene in this example), whether because of a narrower particle distribution or being easier to prepare into a uniform bed. The narrow PSD porous particle achieved nearly identi-cal performance to the core-type particle in the middle veloc-ity region where the A term (bed uniformity) in equation 1 dominates. The core-type particle showed expected perfor-mance advantages over both porous particles at lower veloci-ties (where the B term dominates) and higher velocities (where the C term dominates).

Figure 4b compares a column with narrow-distribution porous silica with two commercial columns that use porous particles with the same spherical shape and average diameter, but broader distribution. As shown, lower hmin values and bet-ter performance for three neutral solutes over the entire f low range were observed for the column with narrow-distribution silica. Table I also confirms that the column with narrow-dis-tribution silica has higher permeability (lower pressure drop or impedance) than those with broader distribution. Figure 5 demonstrates near-equivalency of narrow distribution, sub-2-µm porous and core-type particles using fast 1-min gradient separations on 50 mm × 3.0 mm columns. Although plate height or efficiency cannot be readily compared under gradi-ent conditions, peak width and resolution results indicate that columns are very comparable in performance.

DeStefano (22) compared performance of columns pre-pared with 2.7- and 4.1-µm core-type particles with narrow distribution (D90/10 of about 1.1) to a column made with a 50:50 blend by weight. Results agreed with previously pub-lished data for porous particles that efficient columns could be made with either narrow PSD or broadened PSD core-type silica; however, pressure drop was higher for a column made with blends. Although it may be possible to achieve about the same column efficiency with a broader particle distribution having the same average diameter, separation speed would be more limited because of higher f low resistance. Results are shown in Figure 6 and Table II.

Several other sources of valuable information were used (23–26). Clearly, there is still considerable disagreement about the best particle design for preparing optimum HPLC columns. It should become easier to establish ideal particle properties and optimum column preparation methods as narrower particle distributions become available, computer simulation methods improve, and new bed visualization tech-niques are developed.

Summary and Conclusions

If narrow-PSD particles can be made inexpensively, whether they are porous or core-type, there seem to be advantages to using them. A clear relationship exists between narrower PSD and lower column pressure; however, there is still considerable disagreement about whether narrower distribution leads directly to more uni-form beds and better HPLC column eff iciency. While steady improvements are being made, it is not clear how much performance comes from better particle design or better column preparation techniques. In the absence of any real negatives, particles for HPLC columns are likely to evolve toward narrower distribution. The follow-ing points can be taken from the experiments and data reviewed in this article:t� For the same average particle size and shape, a broader

distribution will generate more f low resistance, presum-ably because smaller particles can fill into gaps between larger ones.

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D I S C O V E R

Why Smart

Scientists Use SAM

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t� With good column preparation tech-niques, broader distribution particles have achieved comparable efficiency in many cases to particles with narrower distribution; however, higher pressure drop will limit those columns to lower

flow velocities and be detrimental to kinetic performance. For the same average particle size and comparable column efficiency, a narrow particle distribution results in significantly lower pressure at the same flow velocity.

t� Verzele noted a small negative impact on performance of broader distribu-tion particles at high flows; while logical, further studies are needed with all particle types to confirm this behavior and establish its importance.

t� New porous silica particles with the same narrow distribution of core-type particles have shown per-formance, measured by hmin, that is better than broader-distribution porous particles and comparable to core-type particles when compared by their reduced plate heights or when gradients are used for separa-tion; having these particles for direct comparison should help establish whether unique core-type particle performance is because of narrow PSD, surface roughness, higher particle density, or something else. Whatever the reason for the high performance of core-type particles, they are widely available and have become well established in methods, so they are here to stay.

t� A major weakness of preparing beds from loose particles is that they can move and settle during use and thereby lose performance; more studies about the impact of particle distribution on column bed stability are needed.

t� Techniques for direct visualization of particle uniformity within the bed will be very valuable in designing ideal particles and standardizing col-umn preparation techniques. Although much is being learned

about designing particles and preparing uniform, efficient particle beds, only part of HPLC column performance can be understood by studying the A term and f low uniformity around par-ticles. Even with a perfect column bed, slow mass transfer processes within the particles and a positive slope for the C term will increase H and reduce over-all column performance, especially at high f low velocities. The C term may be even more difficult to optimize than the A term because we do not know very much about particle pore structure or the stationary-phase environment.

Acknowledgments

Private discussions and unpublished data were provided by Joseph DeStefano

Table II: Comparison of 50 mm × 3.0 mm column performance for single-sized core-type particles and a mixture* (courtesy of Advanced Materials Technology)

Fused-core particles dp90 dp10dp90/dp10

Plate Num-ber (N)†

Plate Height (H)†

Pres-sure (bar)†

Plates per bar

4.1 µm 4.30 3.89 1.11 8590 5.58 34 253

50/50 mix 4.1/2.7 µm 4.26 2.63 1.62 10650 4.69 94 113

2.7 µm 3.02 2.69 1.12 11950 4.19 128 93

* Unpublished data supplied by Advanced Materials Technology.† Measured at optimum �ow (minimum H) in Figure 6. Optimum �ow will be higher for smaller average particle sizes (see Figure 6 for details).

0.2 0.4 0.6 0.8Time (min)

1

1

2

2

3

3

4

4

5

6

6

5

Porous Titan C18, 1.9 �m5800 psi (400 bar) at gradient start

Core-Type C18, 1.7 �m7455 psi (514 bar) at gradient start

1.0 1.2 1.4

0.2 0.4 0.6 0.8Time (min)

1.0 1.2 1.4

Figure 5: Comparison of high-speed liquid chromatography–mass spectrometry (LC–MS) gradients with sub-2-µm monodisperse porous and core-type silica columns. Column dimensions: 5 cm × 3.0 mm; mobile-phase A: 0.1% formic acid in 95:5 water–acetonitrile; mobile-phase B: 0.1% formic acid in 5:95 water–acetonitrile; gradient: 35–60% B in 1 min, then 60% B for 0.5 min; �ow rate: 0.6 mL/min; column tempera-ture: 35 °C; detection: time-of-�ight MS, ESI+, XIC, 100–1000 m/z; injection volume: 2 µL; sample: 300 ng/mL in 97:3 water–methanol. Peaks: 1 = oxazepam glucuronide (463 m/z), 2 = lorazepam glucuronide (497 m/z), 3 = temazepam glucuronide (477 m/z), 4 = oxazepam (287 m/z), 5 = lorazepam (321 m/z), 6 = temazepam (301 m/z). (Unpublished data supplied by Supelco division of Sigma-Aldrich.)


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of Advanced Materials Technology; David Bell, Hugh Cramer, William Campbell, Stacy Squillario, and Wil-liam Betz of the Supelco division of Sigma-Aldrich; Ron Majors, “Column Watch” and “Sample Prep Perspectives” editor for LCGC North America; Wil-liam Barber of Agilent Technologies; and Peter Myers of the University of Liverpool, UK.

Zorbax is a registered trademark of Agilent Technologies; Fused-Core is a registered trademark of Advanced Materials Technology, Inc.; and Titan is a trademark of Sigma-Aldrich Com-pany, LLC.

References

(1) J.H. Knox and M. Saleem, J. Chromatogr.

Sci. 7, 614–622 (1969).

(2) U.D. Neue, HPLC Columns, Theory, Tech-

nology and Practice (Wiley-VCH, Wein-

heim, Germany, 1997).

(3) R.E. Majors, “Directions in HPLC Tech-

nology- Post Verzele,” presented at the

International Symposium on HPLC,

Ghent, Belgium, 2007.

(4) S.A. Schuster, T.J. Waeghe, W.L. Johnson,

J.J. DeStefano, and J.J. Kirkland, “Larger

Superficially Porous Particles,” presented

at the Eastern Analytical Symposium,

Somerset, New Jersey, 2013.

(5) J.H. Knox, J. Chromatogr. A 831, 3–15 (1999).

(6) J.H. Knox, J. Chromatogr. A 960, 7–18

(2002).

(7) K.D. Bartle and P. Meyers, Capillary

Electrochromatography, in RSC Chroma-

tography Monographs, R.M. Smith, Series

Ed. (Royal Society of Chemistry, 2001),

chapter 3.

(8) Horiba Instruments, Inc., A Guidebook to

Particle Size Analysis, www.horiba.com/us/

particle (2012).

(9) C. Dewaele and M. Verzele, J. Chromatogr.

260, 13–21 (1983).

(10) J.J. Kirkland, Anal. Chem. 41, 218–220

(1969).

(11) J.J. Kirkland, Anal. Chem. 64, 1239–1245

(1992).

(12) J.J. Kirkland, J. Chromatogr. A 890, 3–13

(2000).

(13) J.J. Kirkland, J. Chromatogr. Science 38,

535–544 (2000).

(14) J.J. Kirkland, T.J. Langlois, and J.J. DeSte-

fano, Am. Lab. 39, 18–21 (2007).

(15) J.J. DeStefano, T.J. Langlois, and J.J. Kirk-

land, J. Chromatogr. Sci. 46, 254–260

(2008).

(16) F. Gritti, I. Leonardis, D. Schock, P. Ste-

venson, A. Shalliker, and G. Guiochon, J.

Chromatogr. A 1217, 1589–1603 (2010).

(17) D. Cabooter, A. Fanigliulo, G. Bellazzi,

B. Allieri, A Rottigni, and G. Desmet, J.

Chromatogr. A 1217, 7074–7081 (2010).

(18) A. Liekens, J. Billen, R. Sherant, H. Ritchie,

J. Denayer, and G. Desmet, J. Chromatogr.

A 1218, 6654–6662 (2011).

(19) F. Gritti, T. Farkas, J. Heng, and G. Guio-

chon, J. Chromatogr. A 1218, 8209–8221

(2011).

(20) W.E. Barber, B.A. Bidlingmeyer, and X.

Wang, “Interrelations Between Particle

Properties and Chromatographic Perfor-

mance,” presented at the International

Symposium on HPLC, Anaheim, Califor-

nia, 2012.

(21) R.A. Henry, W.R. Betz, W.H. Camp-

bell, G. Parmar, W.K. Way, and P. Ross,

“Development of a New Monodisperse

Porous Silica for UHPLC,” presented at

the International Symposium on HPLC,

Anaheim, California, 2012.

(22) J.J. DeStefano, S.A. Schuster, R.S. Bichl-

meir, and W.L. Johnson, “Particle Size

Considerations in Superficially Porous

Particles,” presented at the Eastern Ana-

lytical Symposium, Somerset, New Jersey,

2013.

(23) A. Daneyko, A. Holtzel, S. Khirevich, and

U. Tallarek, Anal. Chem. 83, 3903–3910

(2011).

(24) S. Bruns, J.P. Grinias, L.E. Blue, J. Jor-

genson, and U. Tallarek, Anal. Chem. 84,

4496–4503 (2012).

(25) K.K. Unger, Porous Silica — Its Properties

and Use as Support in Column Liquid Chro-

matography (Elsevier Scientific Publish-

ing Co., Amsterdam, Oxford, New York,

1979).

(26) R.K. Iler, The Chemistry of Silica (John

Wiley & Sons, Hoboken, New Jersey,

1979).

Richard A. Henryreceived his BS in Chemistry from Juniata College and his PhD in Analytical Chemistry from The Pennsylvania State University. After postdoctoral work at Purdue University, he joined DuPont at the Experimental Station in Wilmington, Delaware and worked with Dr. Jack Kirkland and others in early HPLC column and instrument development. He later became Director of Analytical Laboratories at Penn State University and also founded Keystone Scientific, Inc., to manufacture HPLC column products. Dick remains active teaching short courses on separation technology and as a consultant. Direct correspondence to: [email protected] �

For more information on this topic, please visit


8.0

7.5

7.0

6.5

6.0

5.5

5.0

4.5

4.0

3.5

1 2 3 4 5

2.7 �m

50% 2.7 �m,50% 4.1 �m

4.1 �m

6 7

Pla

te h

eig

ht

(µm

)

Linear mobile phase velocity (mm/s)

Figure 6: Plate height (H) versus velocity (µ) plots for single and mixed core-type particles in 50 mm × 3.0 mm columns. A 50:50 (w/w) mix of 2.7- and 4.1-µm particles produces plate heights intermediate to the single components. The reduced plate heights are about equal. Data are � tted to the Knox equation. (Unpublished data supplied by Advanced Materials Technology.)


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Timothy J. Ward and Karen D. WardMillsaps College, Jackson, Mississippi. Direct correspondence to: [email protected]

Recent Progress in Chiral Stationary Phase Development and Current Chiral Applications

Chiral separations remain a decided area of interest, particularly

in the pharmaceutical and agrochemical fields. Although high

performance liquid chromatography (HPLC) remains a strong

choice for separations because of its robustness, transferability,

and instrument availability, the use of chiral supercritical fluid

chromatography (SFC) continues to expand in analytical and preparative

techniques. Several chiral stationary phases continue to enjoy wide

use because of their broad application in both HPLC and SFC.

Chiral separations continue to be of great interest because of the preva-lence of racemates in markets such

as the pharmaceutical and agrochemical (pesticide) industries. In fact, a review of the importance of pharmaceutical chiral separations in single-enantiomer patent cases was recently published (1), and another review estimates that about 30% of pesticides are chiral with about half of these having multiple chiral features (2). The individual pesticide enantio-mers may exhibit different effects on the environment. Although the separation of enantiomers can be challenging because of their identical physical and chemical properties in an achiral environment, chiral stationary phases (CSPs) have greatly facilitated enantioseparations in high performance liquid chromatogra-phy (HPLC) and supercritical fluid chro-matography (SFC). Research on special-ized separation techniques using novel CSPs, particularly derivatized polysac-charides and cyclodextrins, continues for the resolution of specific individual enantiomers, and chiral separation on commercially available CSPs remains a mature and widely used technique with some new entries to the market. Polysac-charide and macrocyclic glycopeptides CSPs continue to be the most widely used commercial chiral phases, with

cyclodextrins, cyclofructans, π-complex, and protein-based CSPs also finding use. For non-HPLC separations such as gas chromatography (GC) and capillary electrophoresis (CE), cyclodextrins con-tinue to dominate. Enantioseparations of larger, more-complex molecules with multiple chiral centers have increased as biotech continues to grow, meaning that more compounds must be resolved simultaneously, and chiral separations of more-polar molecules are needed, espe-cially for the agrochemical and pharma-ceutical fields.

There are plenty of resources and infor-mation available for performing chiral separations, both from the commercial suppliers of CSPs and in the literature. There are numerous reviews of the widely used CSPs, including recent reviews of cellulose and polysaccharide-based CSPs (3), protein and glycoprotein CSPs (4), macrocyclic antibiotic CSPs (5), cyclo-dextrin CSPs (6,7), and chiral ion- and ligand-exchange CSPs (8). Reflecting the burgeoning interest in SFC chiral separa-tions of pharmaceuticals, several reviews specific to SFC have recently been pub-lished (9–12).

The State of Current CSPs

Chiral separation continues to be the primary technique of choice, with many


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companies seeing an increase of about 20% for both analytical and preparative enantioseparations in their laboratories. The market continues to enjoy growth and maturation as older technologies are replaced by newer and improved technologies. New CSPs continue to be introduced to the market, includ-ing the zwitterionic phases, Chiralpak Zwix(+) and Zwix(-) from Chiral Technologies, a new immobilized crown-ether phase, Chiralpak CR-I also from Chiral Technologies, and an immobilized ovomucoid phase, C18-Ovo-5-120 from Separa-tion Methods Technologies. Companies such as YMC Amer-ica, Inc., Chiral Technologies, Diacel, and Separation Meth-ods Technologies are continuing to expand their offerings of immobilized polysaccharide-derived CSPs because these col-umns offer greater stability, can be used with a wider variety of mobile phases, and are useful in both liquid chromatography (LC) and SFC applications.

One apparent trend in the market is toward the increased use of SFC for chiral separations. The advantages of SFC are the reduced environmental impact and operating costs with increased throughput. Although SFC has traditionally found more use in preparative-scale chiral separations, where the waste reduction and decreased solvent use is attractive to indus-try, these immobilized CSPs are also seeing increased use in analytical applications as well. SFC is currently receiving a lot attention from the pharmaceutical industry for screening and method development of chiral separations (13–15).

Polysaccharide CSPs

As in previous years, chiral separations achieved by polysaccha-ride CSPs in HPLC account for approximately one-third of all HPLC chiral separations in the literature, and most of these sepa-rations are carried out on commercially available columns. The most commonly used phase on cellulose or amylose continues to be 3,5-dimethylphenyl carbamate, which includes immobilized columns such as Chiralpak IA (Chiral Technologies), Chiralpak IB, Lux Cellulose-1 (Phenomenex), and coated phases such as Chiralcel OD-H (Chiral Technologies), Kromasil CelluCoat (Akzo Nobel), and AmyCoat (Akzo Nobel). Because the con-formation of the polymer is influenced by how the stationary phase is attached to the packing material, coated and immobi-lized columns can exhibit different selectivity. Chloro-substi-tuted polysaccharide CSPs have also found much use, including tris(3,5-dichlorophenyl carbamate) (Chiralpak IC), tris(3-chloro-4-methylphenyl carbamate) (Lux Cellulose-2), tris(5-chloro-2-

methylphenyl carbamate) (Chiralpak AY-H), and tris(3-chloro-4-methylphenyl carbamate) (Chiralcel OZ-H). The selectivity of the Chiralpak IC column toward different compounds is dem-onstrated in Figure 1.

A reversed-phase study using tris(chloromethylphenyl carba-mate) derivatives of cellulose and amylose concluded that using these CSPs in screening protocols yields higher success rates in achieving baseline separations with shorter screening times (16). An updated generic separation strategy in normal-phase HPLC was reported using the following commercial CSPs: Lux Cellulose-1, Lux Cellulose-2, Lux Amylose-2, Lux Cellulose-4 and Chiralpak AD-H, Chiralcel OD-H, and Chiralcel OJ-H (17). Reversed-phase screening strategies for HPLC with Chi-ralpak IA, Chiralpak IB, and Chiralpak IC with applications

0 2 4 6

1-Benzoyl-2-tert-butyl-3-methyl-4-imadazoline 1-Phenylethyl-3,5-dinitrobenzoate

8

DevinrolCH

3

CH3

CH3

N

NN

O2N

NO2

O

O

O

O

O

OO

10 12 14 0 2 4 6 8 10 12 14 0 2 4 6 8 10 12 14

Figure 1: The separation of three compounds using a 250 mm × 4.6 mm, 5-µm dp Chiralpak IC column (Chiral Technologies). Mobile phase: 70:30:0.1 (v/v/v) n-hexane–ethyl acetate–diethylamine. Adapted from reference 24.

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compatible with liquid chromatogra-phy–mass spectrometry (LC–MS) were also reported (18). Simplified screening protocols for chiral separations in HPLC and SFC on Chiralpak IA, IB, IC, and ID in reasonable time frames and high success rates were recently published (14).

Cyclodextrin-Based CSPs

Although no new cyclodextrin based CSPS have been brought to market lately, the use of cyclodextrins still accounts for over one-fourth of all publications in chiral HPLC in recent years. The bonded cyclodextrin–based CSPs are popular because of their robustness, wide selectivity, and ability to enantioseparate in the reversed and polar organic phases. The mechanical stability of the cyclodextrin CSPs lends itself to preparative-scale use as well.

Macrocyclic Glycoprotein CSPs

The macrocyclic glycoprotein CSPs con-tinue to fill a broad and useful niche in chi-ral HPLC because of their unique versatility and broad selectivity, and are unsurpassed in the enantioseparation of chiral amino acids. Sigma-Aldrich offers a Chirobiotic method development kit containing V2, T, R, and TAG columns for screening in polar ionic, polar organic, reversed-phase, and normal-phase modes. A review of macro-

cyclic glycopeptide-based CSPs in HPLC methods for amino acid enantiomers and related analogues was published in 2010 (19) and recently updated (5).

Cyclofructan CSPs

The recently released cyclofructan-based Larihc CSPs from AZYP, a new sup-plier of novel chiral and achiral phases for HPLC, hydrophilic-interaction chro-matography (HILIC) and SFC, also available from Supelco/Sigma-Aldrich, continue to have great application in chiral HPLC separations. (Larihc is “chi-ral” spelled backwards.) These columns include Larihc CF6-P, considered the

“king column” for separation of racemic primary amines, Larihc CF6-RN, which separates nonprimary amines, and Lar-ihc CF7-DMP, the only commercialized cyclofructan 7 column, which shows complementary enantioselectivity to the CF6-RN CSP.

Larihc CF6-P is the only column that separates primary amines in a non-aqueous solvent using the polar organic mode. The Larihc CF6-P was reported to be useful in the separation of chiral illicit drugs and controlled substances, with the other Larihc CSPs also yielding enantioseparations (20). A comparison of separations of 46 chiral reagents to deter-

mine enantiomeric purity using Larihc, Cyclobond (Supelco/Sigma-Aldrich), and Chirobiotic CSPs was recently reported, with this being the first use of Larihc CSP for this purpose (21). Recent reports indicate that the Larihc CF6-P CSP is broadly applicable to the separa-tion of nonamine containing racemates as well (20–22).

All Larihc CSPs are reported to work well in SFC because of their use in the normal-phase or polar organic phase modes. A comparison of dimethylphenyl carbamate cyclofructan 7 CSP use in SFC and HPLC was recently made (23) and the retention and enantiodiscrimi-nation properties and the effect of differ-ent SFC modifiers was reported.

Chiral Separations Today

Although the chiral market is maturing and stable, perhaps the biggest need in the field remains an increased overall under-standing of chiral methodologies and more practical training. Because infor-mation is largely supplied by vendors to newcomers to chiral HPLC, novices can sometimes stumble around for a while wading through vendor literature and advice. In addition to an understand-able sales objective, vendors occasionally misunderstand the customers’ end-to-end chiral operation, so promised increases are sometimes not realized at the labo-ratory level. Furthermore, increased per-formance is also sometimes not realized because of low-tech reasons that may be related more to laboratory layout and operation, rather than the lack of latest technology and equipment. Newcomers should make use of the short courses on chiral HPLC that give a wealth of unbi-ased information to the participants and are offered at major conferences such as Eastern Analytical Symposium (EAS), The Pittsburgh Conference (Pittcon), and Chirality.

Increased selectivity remains a chal-lenge in chiral separations. With increased selectivity, loading can be increased in preparative chiral separations, which will provide the greatest cost savings. With significant cost savings in the separation process, companies can shift from using chiral selective synthesis for chiral purifi-cation to purification by chromatography. Although HPLC is used for preparative-scale separations, SFC continues to gain

O

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Bendro�umethiazide

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NH2

HN

O O O O

SS

NH

CF3

Figure 2: Examples of separations of pharmaceutical compounds on CF6 and CF7 CSPs. Adapted from reference 25.


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ground as the most cost-effective way to purify enough material for further studies.

The market is maturing, but technol-ogy has not moved forward in any revolu-tionary way. Chiral HPLC still lacks (and may always lack) a CSP that can operate in all modes for enantioseparation of mol-ecules from low to high polarity.

Acknowledgments

We gratefully thank Daniel W. Armstrong, Elena Eksteen, Hafeez Fatunmbi, J.T. Lee, and Gary Yanik for their suggestions in the preparation of this manuscript.

References

(1) C. Weekes, Drugs Pharm. Sci. 211, 304–311

(2012).

(2) E. Ulrich, C. Morrison, M. Goldsmith, and

W. Foreman, Rev. Environ. Contam. Toxicol.

217, 1–74 (2012).

(3) J. Shen and Y. Okamoto, Compr. Chirality 8,

200–226 (2012).

(4) J. Haginaka, Compr. Chirality 8, 153–176

(2012).

(5) I. Ilisz, Z. Pataj, A. Aranyi, and A. Peter, Sep.

Purif. Rev. 41, 207–249 (2012).

(6) Y. Xiao, S. Ng, T. Tan, and Y. Wang, J. Chro-

matogr. A 1269, 52–68 (2012).

(7) X. Zhang, Y. Zhang, and D.W. Armstrong,

Compr. Chirality 8, 177–199 (2012).

(8) B. Natalini and R. Sardella, Compr. Chirality

8, 115–152 (2012).

(9) R. Wang, T. Ong, S. Ng, and W. Tang,

Trends Anal. Chem. 37, 83–100 (2012).

(10) K. De Klerck, D. Mangelings, and Y. Vander

Heyden, J. Pharm. Biomed. Anal. 69, 77–92

(2012).

(11) C. West, Curr. Anal. Chem. 10(1), 99–120

(2014).

(12) K. De Klerck, Y. Vander Heyden, and D.

Mangelings, J. Chromatogr. A 1328, 85–97

(2014).

(13) L. Kott, Am. Pharm. Rev. 16(1), 5/1–5/8

(2013).

(14) K. De Klerck, C. Tistaert, D. Mangelings,

and Y. Vander Heyden, J. Supercrit. Fluids

80, 59–59 (2013).

(15) W. Schafer, T. Chandrasekaran, Z. Pirzada,

C. Zhang, G. Chaowei, B. Xiaoyi, and R.

Mirlinda, Chirality 25(11), 799–804 (2013).

(16) L. Peng, S. Jayapalan, B. Chankvetadze, and

T. Farkas, J. Chromatogr. A 1217(44), 6942–

6955 (2010).

(17) A.A. Younes, D. Mangelings, and Y. Vander

Heyden, J. Pharm. Biomed. Anal. 56(3),

521–537 (2011).

(18) T. Zhang, D. Nguyen, and P. Franco,

J. Chromatogr. A 1217(7), 1048–1055

(2010).

(19) I. Ilisz, Z. Pataj, and A. Peter, Macrocyclic

Chem. 129–157 (2010).

(20) N. Padivitage, E. Dodbiba, Z. Breitbach, and

D.W. Armstrong, “Enantiomeric separations

of illicit drugs and controlled substances

using cyclofructan-based (LARIHC) and

cyclobond I 2000 RSP HPLC chiral station-

ary phases,” Drug Test. Anal. doi:10.1002/

dta.1534 (2013).

(21) H. Qiu, N. Padivitage, L. Frink, and D.W.

Armstrong, Tetrahedron: Asymmetry, 24(18),

1134–1141 (2013).

(22) J. Smuts, X. Hao, Z. Han, C. Parpia, M.

Krische, and D.W. Armstrong, Anal. Chem.

86(2), 1282–1290 (2014).

(23) J. Vozka, K. Kalikova, C. Roussel, D.W.

Armstrong, and E. Tesarova, J. Sep. Sci.

36(11), 1711–1719 (2013).

(24) http://immobilizedchiralcolumns.com/csp-

robustness.

(25) http://www.sigmaaldrich.com/etc/medialib/

docs/Supelco/Posters/1/daw-chiral-010711.

Par.0001.File.tmp/daw-chiral-010711.pdf.

(26) http://www.sigmaaldrich.com/etc/medialib/

docs/Supelco/Posters/1/daw-chiral-010711.

Par.0001.File.tmp/daw-chiral-010711.pdf.



Timothy J. Ward is a professor of chemistry and associate dean of sciences at Millsaps College (Jackson, Mississippi). Ward received his BS degree from the University of Florida and his PhD from Texas Tech University. Dr. Ward served as chair of the International Symposium on Chirality in July 2007, in San Diego, California. His research interests include chiral separations, the development of analytical LC and CE methods, and their application to pharmaceutical and archaeological analysis.

Karen D. Ward is an instructor at Millsaps College. She received her BS degree from Texas A&M University and her MS from Texas Tech University. Ms. Ward previously worked in the pharmaceutical industry at the Analytical Environmental Research Division at Syntex Pharmaceuticals in Palo Alto, California.Direct correspondence to: [email protected] �

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Figure 3: Chromatograms showing chiral separations in normal-phase, polar organic phase, and reversed-phase modes on CF6-RN CSP. Adapted from reference 26.


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C. David CarrBioanalytical Technologies. Direct cor-respondence to: [email protected]

The Role of Chromatography in the Characterization and Analysis of Protein Therapeutic Drugs

Chromatography has taken a prominent place in the characterization and

analysis of protein therapeutic drugs and today it plays a critical role in

the biotechnology laboratory. Although reversed-phase chromatography

is the foremost chromatography technique used for this purpose,

other techniques such as ion-exchange, size-exclusion, normal-phase,

hydrophilic-interaction, and hydrophobic-interaction chromatography

play very specific roles in characterizing and analyzing protein drugs.

Chromatography has long been an essential tool for protein purification. Dextran-based ion-

exchange and size-exclusion columns have been instrumental in protein puri-fication and characterization for decades. With the development of recombinant DNA proteins for therapeutic purposes there has been a need for careful, detailed characterization of these proteins, the changes and modifications that might occur, as well as sensitive determination of changes that actually occur. Chroma-tography has taken its place as a premier technology in both the characterization and the analysis of recombinant protein therapeutic drugs (1).

Protein Modifications

and Their Effect on

Protein Therapeutic Drugs

The biological activity and, therefore, the therapeutic efficacy of protein drugs depend on the exact composition and three-dimensional structure of the pro-tein. Any change to a protein (modifica-tion) may affect its efficacy as a thera-peutic agent, although efficacy may not be affected by some protein modifica-tions. The effect of a given modification depends on the protein and the nature and location of the modification. Dur-ing development of a protein therapeutic

drug it is necessary to fully characterize the protein, define what possible modi-fications may occur, define those modi-fications that do affect efficacy (critical quality attributes), and develop methods to analyze such modifications. Changes occur to proteins from the beginning of their assembly. Some changes may be needed for the protein to be properly formed into an active tertiary struc-ture such as glycosylation and disulfide bonds, which are critical in maintain-ing the proper structure. Glycosylation attaches sugar chains to either aspara-gine or serine residues. These sugar chains affect protein folding and assist in maintaining correct tertiary protein structure. Glycosylation structure is complicated and is affected by the cell lines used for protein expression as well as other aspects of the expression system. It is vital to characterize and monitor glycosylation patterns. Disulfide bonds formed between cysteine residues are essential for formation and maintenance of proper tertiary structure. These must be characterized as to their locations in the primary structure of a protein and also must be monitored to ensure that a protein remains active.

Other modifications may affect pro-tein binding and reduce activity such as chemical deamidation — the conversion


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of an asparagine residue into an aspartic or isoaspartic acid residue under condi-tions of high temperature or high pH

— and oxidation of methionine residues under chemical oxidative conditions to methione sulfoxide. These two modifi-cations may affect activity if found near catalytic or binding sites, but may have no effect if found distant from sites of activity.

Some modifications are engineered into proteins to form more effective drugs. The addition of polyethylene gly-col to proteins — pegylation — usually extends the duration of protein activ-ity in the blood stream several times by reducing the rate at which the pro-

tein is removed from the blood stream. Monoclonal antibodies (mAbs) that bind to specific receptors on the surface of tumor cells can be modified by the addition of highly toxic drugs to the monoclonal antibody, creating a highly specific, effective reagent that targets and kills specific tumor cells. Engi-neered modifications must be analyzed and monitored.

Proteins may lose tertiary structure by variations in pH, temperature, or the presence of certain reagents. Such “dena-tured” proteins lose biological activity and pharmacological potency. Proteins may also form “aggregates,” where mul-tiple proteins self-associate into larger

complexes that usually reduce activity and may engender immune responses in patients.

Reversed-Phase HPLC

The most powerful and useful chroma-tography technique used for analysis and characterization of proteins drugs is reversed-phase high performance liquid chromatography (HPLC). This technique is most commonly used to separate, identify, and measure peptides that result from the enzymatic diges-tion of the therapeutic protein (peptide map). Although a single modification on a large, many kilodalton protein may cause enough of a change in the retention of the protein in reversed-phase HPLC to differentiate the modi-fied protein from the native protein, often the change is too small to cause a measurable change in retention. By digesting a protein into smaller pieces

— typically from a very few amino acids to 20–40 amino acids — the modifica-tion now makes a much larger change in the retention of the modified peptide relative to the native peptide than for the intact protein, thus allowing easier detection and quantitation of the modi-fication. Figure 1 shows the effect on peptide retention of the addition of a methionine residue to human growth hormone, which occurs when the pro-tein is expressed in E. coli bacteria. Such peptide maps are the basis of many chromatography methods for both char-acterization and analysis of therapeutic proteins. Among protein modifications that are easily detected and measured by peptide maps are deamidation of an asparagine residue, oxidation of a methionine residue, and conversion of an N-terminal glutamine residue to cyclized glutamate. These modifica-tions may diminish the efficacy of a therapeutic protein or otherwise affect its potency as a therapeutic agent. Other protein characteristics that can be mon-itored by peptide maps include disulfide bonds between two cysteine residues and exact locations of glycosylation. Reversed-phase HPLC is widely used in the analysis and characterization of therapeutic proteins because the list of modifications that can be characterized, identified, or quantified by this technique is very long.

Peptide map of hGH expressed in E.coli

Peptide map of native hGH

N-terminal peptide

N-terminal peptide

with added Met

Figure 1: Effect of the addition of a methionine residue to human growth hormone be-cause of expression in E. coli on the relative retention of a peptide in a peptide map. Met is somewhat hydrophobic so the peptide containing Met is retained longer than the (native) peptide absent the Met residue. Adapted from reference 1.

Figure 2: Polypeptides are large molecules that adsorb strongly to the hydrophobic surface and desorb only at a very speci�c solvent strength. A portion of the polypeptide adsorbs to the surface (stationary phase), and some of the molecule remains in contact with the mobile phase. See reference 3.


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The Separation of Polypeptides

by Reversed-Phase HPLC

Although reversed-phase HPLC has been known and practiced for several decades, initially it was considered impractical for separating large molecules such as pro-teins or even peptides. Early attempts at protein or peptide separations by reversed-phase HPLC resulted in poor peak shapes and separations. The intro-duction of larger-pore silica (average pore size: 150–300 Å) for chromatogra-phy columns in the early 1980s provided

the first really high resolution of large polypeptides (2,3). This discovery coin-cided with the early days of recombinant DNA protein development and the two grew hand in hand. The use of large pore silica opened the door to rigorous development of optimal conditions for polypeptide separations and, today, such optimized conditions coupled with high quality reversed-phase HPLC separation columns result in separations that are highly effective in characterizing and monitoring therapeutic drugs.

Larger pores allow the entrance of the larger protein and peptide molecules into the interior of the pores where the bulk of the adsorptive surface causing separation is found. The relatively large polypeptide then adsorbs to the surface of the modified, hydrophobic surface (Figure 2). Large polypeptides are more strongly bound to the hydrophobic sur-face than smaller molecules and do not desorb from the surface until a stronger (for example, more hydrophobic solvent) enters the column and pores. Once desorbed, the polypeptide appears to interact minimally with the surface of the silica during its transport down the column. The amount of separation is essentially determined by the strength of that initial adsorption, which in turn is determined by the size and hydrophobic-ity of the portion of the polypeptide that actually interacts with the hydrophobic surface. This portion of the molecule is called the hydrophobic foot (4) and is the key to polypeptide separations. Because of the large size of the polypeptide mol-ecule, only a portion will interact with and adsorb to the hydrophobic surface. A portion of the polypeptide will be in contact with the mobile phase. Even very small changes to the hydrophobic foot cause a change in the retention of molecules in reversed-phase HPLC resulting in separation. This accounts for the ability of reversed-phase HPLC to separate proteins or peptides that dif-fer in even very small ways from a native or reference protein or peptide. In the separation of two insulin variants shown in Figure 3, a difference of only a single methyl group (rabbit insulin has a threo-nine where human insulin has a serine, a difference of a single methyl group) which represents a difference of 15 MW in a protein of about 5700 MW, and yet reversed-phase HPLC is able to separate these two proteins (5). The ability of reversed-phase HPLC to separate such very similar polypeptides is at the heart of its use in the characterization and analysis of protein therapeutic drugs.

Conditions for

Polypeptide Separations

by Reversed-Phase HPLC

The separation column is based on small, porous silica particles of 1.7–5 µm diameter. Most protein therapeutic

0 5 10 15 20 25

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Rabbit insulin(threonine)

Human insulin(serine)

Figure 3: The ability of reversed-phase HPLC to separate very similar polypeptides is illus-trated by the separation of two insulin variants (human and rabbit) that differ by a single amino acid, which in turn differ by a single methyl group. Adapted from reference 2.

Table I: Chromatography techniques commonly used for protein modification analysis

Modification/Degradation Applicable Chromatography Techniques

Deamidation Reversed-phase HPLC peptide mapping, ion exchange

Oxidation Reversed-phase HPLC peptide mapping

Glycan analysis Reversed-phase HPLC, normal phase, HPAEC, HILIC

Mistranslation Reversed-phase HPLC peptide mapping

Aggregation Size exclusion

Denaturation Size exclusion

Disul�de bonds Reversed-phase HPLC peptide mapping

Charge variants Ion exchange

ConjugationSize exclusion, reversed-phase HPLC peptide mapping, HIC, ion exchange

HPAEC = High-pH anion-exchange chromatographyHILIC = Hydrophilic-interaction chromatographyHIC = Hydrophobic-interaction chromatography


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drug separations use 5-µm silica par-ticles, however, smaller particle col-umns are increasing in usage because of shorter analysis times and, sometimes, higher resolution (6) for separation of mAbs using superficially porous silica particles. The best pore size for poly-peptides is normally between 150 Å and 300 Å, although very small peptides, and sometimes enzymatic digests, can be satisfactorily separated using col-umns of 100-Å pore size silica. The polar surface of the silica particles is modified by short (4–18 carbon atoms) aliphatic organic chains. Longer chain groups (for example, 18 carbons) are usually best to separate small peptides and protease digests of proteins while smaller hydrophobic groups (such as four carbons) are best for retention and

separation of larger peptides (>15–20 residues), proteins, or hydrophobic polypeptides.

The mobile phase consists of water, an organic solvent (nearly always acetoni-trile), and an ion-pair reagent. Although acetonitrile is the most commonly used organic solvent when separating poly-peptides, adding a small (1–5%) per-centage of isopropanol can be an aid in the separation of hydrophobic polypep-tides or when sample recovery is low.

The ion-pair reagent used is almost always trif luoroacetic acid, which evap-orates well, is relatively UV transparent, and results in sharp polypeptide peaks and good resolution. Trif luoroacetic acid is usually added to the mobile phase at a concentration of about 0.1%. With peptide maps, the concentration

of the trif luoroacetic acid may affect resolution between peptide pairs to a small extent and trif luoroacetic acid concentrations of 0.1–0.5% have been used to achieve slightly better resolu-tion of critical pairs of peptides. When using electrospray mass spectrometry (MS) with reversed-phase HPLC, the use of trif luoroacetic acid results in a much lower signal than obtained in its absence. The signal loss depends on the concentration of trif luoroacetic acid, and low concentrations of 0.01–0.05% result in relatively little signal loss; however, resolution and peak shape may suffer at the lower concentrations of trif luoroacetic acid. It has been shown, however, that very low concentrations of trif luoroacetic acid result in good peak shape and resolution if very high purity silica (Type B) columns are used. Con-sequently, many manufacturers offer very high purity columns specifically for use with trif luoroacetic acid and reversed-phase HPLC–MS. Formic acid has also been used by some polypeptide chromatographers to avoid the signal loss in MS when using trif luoroacetic acid, although some chromatographers have found decreased resolution with formic acid, so it is usually reserved for MS applications. Other ion-pair reagents such as heptaf luorobutyric or pentaf luoroproprionic acid have been used on occasion, particularly for more-hydrophobic polypeptides.

Gradient elution is always used in polypeptide separations. The organic solvent concentration is low (0–5%) at the beginning of the separation and is increased slowly over the course of the separation. Because of the adsorp-tion mechanism of polypeptides on the hydrophobic surface, very small changes in organic solvent concen-tration affect the retention of a poly-peptide, so very small changes in the organic solvent concentration (gradient slope) can have a significant effect on retention and resolution. Shallow gra-dient slopes of 0.1–0.5% are commonly used, resulting in separation times of 30 min to as high as 180 min for sepa-rating complex mixtures of peptides, such as enzymatic digests of very large proteins. Occasionally, the gradient slope will affect the resolution of pep-tide pairs in unexpected ways. Although

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Figure 4: Slight differences in the way that peptides adsorb onto the reversed-phase HPLC surface results in variations in relative retention of some peptides at (a) 20 °C, (b) 40 °C, and (c) 60 °C. This leads to changes in resolution between some peptide pairs in a reversed-phase HPLC peptide map. In this example of the peptide map of human growth hormone, peptides 11, 12, and 13 are poorly separated at 20 °C, are much better separated at 40 °C, but are coeluted at 60 °C. Peptide 15 is eluted before peptide 14 at 20 °C, slightly later at 40 °C, and signi�cantly later at 60 °C. Temperature often has a signi�cant effect on peptide resolution and must be optimized when developing a peptide map separation. Adapted from reference 7.


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reductions in gradient slope (resulting in increased elution times) generally lead to increased resolution, because of differences in the thermodynamics of adsorption of some peptides, the resolu-tion of some peptide pairs may decrease (7) with decreasing gradient slope. Gra-dient slope, then, must be carefully optimized to achieve optimal resolution of the peptides in a protease digest in the best time.

The column temperature also affects the resolution of some peptide pairs because of differences in the thermody-namics of adsorption of some peptides (Figure 4) (8). Because temperature can affect the relative retention of pep-tides, several temperatures should be tried when developing a peptide map separation with careful monitoring of the effect of temperature changes. The optimized column temperature is a very important element in establishing an analysis method and its validation. High temperatures often improve peak shape and resolution when analyzing large polypeptides such as mAbs (9).

Detection of polypeptides in reversed-phase HPLC is normally done at or near 214 nm. This wavelength offers opti-mal detection of polypeptides. During a typical gradient to separate peptides, the baseline may rise because of a shift in adsorption at this wavelength by tri-f luoroacetic acid as the dielectric con-stant of the solvent changes. This can

be compensated by reducing the con-centration of trif luoroacetic acid in the organic solvent by 10–15% relative to the aqueous solvent.

Ion-Exchange Chromatography

Ion-exchange chromatography has long been a useful tool in the purification of proteins and remains a mainstay in the purification of therapeutic protein drugs. Ion-exchange chromatography does not have the resolving power of reversed-phase chromatography, how-ever, and is usually not able to sepa-rate small differences between protein molecules. Despite this, because ion-exchange chromatography separates by a different mechanism than reversed-phase HPLC — it separates proteins by charge rather than hydrophobicity — it can perform certain separations better than reversed-phase HPLC. Proteins bind strongly to the ionized surface of ion-exchange materials by the opposite charges found on the protein. Cation-exchange chromatography, with par-ticles containing negative charges (sul-fonic or carboxylic acid groups), binds a protein by the positive charges of lysine and asparagine (also the amino termi-nus). Anion-exchange chromatography uses particles with positive charges and binds the negatively charged amino acids aspartic and glutamic acids (as well as the carboxy terminus of a pro-tein). Although ion-exchange chroma-

tography lacks the high resolving power of reversed-phase HPLC, it has found use in monitoring modifications to pro-teins that change the overall charge on a protein (called charge variants). This includes modifications such as deami-dation, which adds a negative charge to a protein, converting the neutral aspara-gine to a negatively charged aspartic (or isoaspartic) acid group. Other charge variants are formed when lysine is enzy-matically removed from the carboxy ter-minus of an antibody heavy chain; when N-terminal glutamine cyclizes with the amino terminus, thus losing a positive charge; when molecules such as poly-ethylene glycol are added to primary amines in a protein, thus losing a posi-tive charge; or when acidic sugars (such as sialic acid) are added to the ends of glycan chains. These charge variants can be effectively monitored using ion-exchange chromatography (10). Condi-tions are simple; the ion exchange mate-rials have charged (negative or positive) groups on their surface to which the proteins bind. Elution is accomplished with a simple linear increase in salt — typically sodium chloride — concen-trations in the mobile phase. Gradients from 0–2 M salt suffice to elute almost all proteins. Separation and resolution of proteins is very sensitive to the mobile-phase pH and the optimum pH must be empirically determined for each protein.

The use of ion-exchange and reversed-phase HPLC in multidimensional LC × LC techniques combines the strengths of the two different mechanisms and results in very high resolution separa-tions. For example, the use of a cation-exchange column as the first stage will separate peptides on the basis of charge and individual peptides with similar charge can be further separated by using a reversed-phase second column. The operation can be run with off-line col-lection of the eff luents from column 1 with manual reinjection onto column 2. Some workers prefer on-line coupling of the two modes by the use of high-pressure switching valves after column 1 to direct the effluent to column 2 and allow automation of the entire operation.

Size-Exclusion Chromatography

Another form of chromatography long used in protein purification is size-exclusion

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Figure 5: Glycans are very polar molecules and can be separated and analyzed by hydrophil-ic interaction chromatography. This chromatogram shows the separation of glycans present on the protein bovine fetuin. Data courtesy of Waters Associates.


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chromatography (SEC), sometimes called gel f iltration when using an aqueous mobile phase. SEC separates molecules, particularly proteins, by differences in three-dimensional size, including water bound to the protein (hydrodynamic volume). Although the resolution between proteins of differ-ent sizes is not very good, size exclusion has found an important place in pro-tein therapeutic drug characterization and analysis by separating monomeric proteins from aggregates, a crucial fac-tor in protein analysis. Size exclusion is also a technique useful in monitoring denaturation if that occurs. Columns with different fractionation ranges can be obtained and the mobile phase is designed to simply solubilize the pro-tein very well. The use of SEC in the characterization of proteins and their aggregates is covered elsewhere in this issue (11).

Chromatographic

Analysis of Glycosylation

The nature and structure of sugar chains attached to proteins (glycosyl-ation) must be regularly monitored in protein therapeutic drug batches. This ensures reproducibility of the produc-tion process and minimizes any effects that changes in glycosylation may have on protein activity and therefore drug efficacy. For many years the technique of high-pH anion-exchange chroma-tography (HPAEC) has been used to monitor glycosylation. In recent years other chromatographic techniques and nonchromatography techniques (such as capillary electrophoresis and MS) have been developed to provide better or more sensitive analysis of glycosyl-ation patterns. In HPAEC, the mobile-phase pH is adjusted to a pH of 11–13, thus ionizing the hydroxyl groups on the sugars. These are then separated by anion-exchange chromatography using special instrumentation designed to operate under these corrosive condi-tions. Because sugars do not contain UV chromaphores, detection is done by electrochemical means. To avoid the corrosive conditions of HPAEC, other chromatography techniques have been developed for glycan analysis. In most cases, these techniques require the use of sugar derivatization with f luorescent

molecules, such as anthranilic acid, to facilitate detection. Derivatized glycans can be detected with high sensitivity by f luoresence and can be separated by a variety of chromatographic tech-niques. Reversed-phase chromatogra-phy of derivatized glycans is effective at separating and quantitating glycans (12). Normal-phase chromatography is also effective at separating glycans (13). Normal-phase chromatography relies on the hydrogen bonding of the polar sugar hydroxyl groups to amine groups on the surface of the chromatographic particles for separation. Reversed-phase HPLC and normal-phase techniques have the advantage of being able to be coupled with MS, offering additional information on the nature of the gly-cans. A more recent addition to the chromatographic lineup for glycan anal-ysis is hydrophilic-interaction chroma-tography (HILIC). This technique uses polar particles (often bare silica) with a mobile phase containing high concen-trations of acetonitrile, with relatively low concentrations of water contain-ing salt. The aqueous solution adsorbs onto the particle surface forming a layer of very hydrophilic liquid. Polar com-pounds such as glycans partition from the relatively organic mobile phase into the relative hydrophilic phase adsorbed to the particle surface. Elution is usu-ally by means of an increase in the water content of the mobile phase and a subsequent decrease in the organic phase over time. Glycans, being very polar molecules, separate very well by this technique and effort has gone into developing a practical framework by which to identify and characterize the glycans that are separated by small-par-ticle HILIC columns (Figure 5).

Conclusion

Chromatography has taken a promi-nent place in the characterization and analysis of protein therapeutic drugs and today plays a critical role in the biotechnology laboratory. Although reversed-phase is the foremost chro-matography technique used for this purpose, other techniques such as ion exchange, size exclusion, normal phase, HILIC, and HIC play very specific roles in characterizing and analyzing protein drugs.

References

(1) R.L. Garnick, N.J. Solli, and P.A. Papa,

Anal. Chem. 60(23), 2546–2557 (1988).

(2) J.D. Pearson, N.T. Lin, and F. Regnier,

Anal. Biochem. 124(1), 217–30 (1982).

(3) J. Rivier et.al., J. Chromatogr. 268, 1121

(1983).

(4) X. Geng and F. Regnier, J. Chromatogr.

296, 15–30 (1984).

(5) J. Rivier and R. McClintock, J. Chrom.

268, 112–119 (1983).

(6) S. Fekete, S. Rudaz, J. Fekete, and D.

Guillarme, J. Pharm. Biomed. Anal. 70,

158–168 (2012).

(7) R.C. Chloupek, W.S. Hancock, and L.R.

Snyder, J. Chromatogr. 594, 65–73 (1992).

(8) W.S. Hancock, R.C. Chloupek, J.J. Kirk-

land, and L.R. Snyder, J. Chromatogr. 686,

31–43 (1994).

(9) S. Fekete, S. Rudaz, J-L. Veuyhey, and

D. Guillarme, J. Sep. Sci. 35, 3113–3123

(2012).

(10) Y. Lyubarskaya, D. House, J. Woodard,

and R. Mhatre, Anal. Biochem. 348(1),

24–39 (2005).

(11) L. Lloyd, LCGC North Am. 32(s4), 30–35

(2014).

(12) X. Chen, Y.D. Liu, and G. Flynn, Glycobi-

ology 19(3), 240–249 (2009).

(13) K.R. Anumula and S.T. Dhume, Glycobiology

8(7), 685–694 (1998).



C. David Carris with Bioanalytical Technologies. He has been involved with HPLC for more than 40 years. He has worked with the biotechnology industry for many years in the characterization and analysis of protein therapeutics and is the author of the booklet “The Handbook of Analysis and Purification of Proteins and Peptides by Reversed-Phase HPLC.” For the past 16 years he has taught the class The Analysis and Characterization of Protein Therapeutic Drugs for Bioanalytical Technologies (www.bioanalyticaltech.com) to scientists from most of the major biotechnology firms and many smaller biotech companies.Direct correspondence to: [email protected] �


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Linda LloydAgilent Technologies, Church Stretton, Shropshire, UK. Direct correspon-dence to: [email protected]

Size-Exclusion Chromatography of Protein Aggregation in Biopharmaceutical Development and Production

New biological entities, protein-based pharmaceuticals, are now

routinely obtained by genetic engineering with host cells that are

mostly mammalian and microbial. It is essential that robust analytical

methods are developed to identify and monitor aggregation and

accurately quantify the aggregate content of a biopharmaceutical

preparation. Size-exclusion chromatography is an important

technique in biopharmaceutical characterization, and this article

discusses its use for soluble aggregation analysis and quantitation.

New biological entities, protein-based pharmaceuticals, are now routinely obtained by genetic engineering

with host cells that are mostly mamma-lian and microbial. The cellular processes are complex. Often the resultant recombi-nant proteins are unstable and aggregate or do not adopt the native conformation that imparts the required biological activity. The subsequent multistep purification procedure subjects the target protein to numerous changes in its environment with an associ-ated risk of further conformational changes and increased levels of aggregation, visible precipitation, and invisible soluble aggregates. The impact of aggregation on the process economics, efficacy, and immunogenicity of a biopharmaceutical are considerable, and so reliable and accurate methods of analysis and quantitation that can be applied to the vari-ous scenarios encountered in development and production are required.

It is therefore essential that robust analyti-cal methods are developed to identify and monitor aggregation and accurately quantify the aggregate content of a biopharmaceutical preparation. One technique that is used for soluble aggregation analysis and quantitation is size-exclusion chromatography (SEC).

Aggregation

Protein aggregation can impact both the economic viability of a biopharmaceuti-cal product and its efficacy. The reduction in the economic viability of the process is seen through a reduction in product yield or decreased bioactivity of the product. An increase in the level of aggregation can increase the immunogenicity of the final product because the recipient’s immune system may recognize the protein complex as nonself and trigger an antigenic response.

At the molecular level, the formation of protein aggregates is complex, but it is accepted that as part of the mechanism of formation the protein must at some point lose its three-dimensional structure to interact with other protein molecules. The mechanism of interaction of unfolded pro-teins can result in the formation of irrevers-ible aggregation, but if there was minimal disruption to the three-dimensional struc-ture then aggregation may be reversible. In the worst case, the proteins can irreversibly denature and the three-dimensional struc-ture and, hence, bioactivity, is lost. In this case the protein no longer functions as a biopharmaceutical, efficacy is reduced, and the process yield is decreased (1).


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Novel Chemistry, Improved Selectivity

www.imtaktusa.com [email protected]

No pre- or post- labeling required

LC-MS analysis for 55 amino acids in 10 minutes

Intrada Amino Acid, 50 x 3 mmA: ACN / THF / 25mM Ammonium formate / Formic acid = 9 / 75 / 16 / 0.3B: ACN / 100mM Ammonium formate = 20 / 80Gradient Conditions:0 %B (0-2.5min)0-17 %B (2.5-6.5 min)100 %B (6.5-10 min)0.6 mL/min (6MPa), 35 °C, 5 �L (1�mol/mL)ESI (SIM, positive)

INTRODUCING:

Intrada Amino Acid HPLC Column


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Aggregation in the Develop-

ment and Production Process

Protein aggregation is encountered dur-ing various stages of biopharmaceutical manufacture and storage because it is one reaction to changes in the protein environment or forces exacted by the environment — chemical and mechani-cal stress (2). Examples include changes in solution interfaces, interactions with surfaces or solids, mechanical stresses because of f low or agitation (3), and changes in temperature.

Upstream — Cell CultureThe expression of biopharmaceuticals is mainly carried out using a fed-batch process with a time line of multiple days — typically 10–14. During this first stage in the manu-facturing process, there will be numer-ous changes in the environment, referred to as cell culture media, as nutrients are consumed and replenished and the pH is adjusted to maintain optimum expression. During the optimization of the upstream process, aggregation is monitored and con-ditions that promote aggregation are identi-

fied. This enables the cell culture to be run at a pH and salt concentration in which the protein is most stable and, where necessary, molecules can be added that assist with sta-bilization. The conditions for maximum stability or minimum aggregation identified here can also be used as the basis for storing the protein substance and biopharmaceuti-cal formulation. During development and optimization of the cell culture, a high num-bers of samples are produced, so methods of analysis must be able to deliver results in a short time frame. By using a liquid chroma-tography (LC) system configuration with shorter columns and higher flow rates, SEC can meet the sample throughput needs.

Downstream — PurificationAlthough there is a need for fast separations during the development of downstream pro-cesses, methods capable of delivering accu-rate quantitation are also needed because they will form the basis of release criteria. SEC methods with increased resolution are needed to achieve robust quantitation.

The capture of the target protein from the cell culture and its subsequent puri-fication is a multistep process that often includes one step specifically designed for the removal of protein aggregates — ion-exchange or size-exclusion purification. However, the downstream purification steps can also increase the concentration of aggregates through changes in the protein microenvironment, concentration, pH, and salt concentration. The chromatographic purification conditions are chosen so as to minimize aggregate formation, including mobile-phase selection and reducing the on-column and elution protein concentra-tion, but in-process testing is needed at each stage of the purification to ensure that the level of aggregation is as expected.

Formulation StudiesUnlike small-molecule drugs, a biothera-peutic is administered as a formulation and, therefore, its stability, including aggregation, must be determined in formulations and conditions that match as closely as possible those in which the drug is applied in vivo. As part of the early stage in formulation development, the environmental effects required to obtain acceptable product shelf life and conditions that control aggregate formulation must be identified. Variables that must be considered include protein concentration, temperature, ionic strength,

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Figure 2: Separation of the BioRad protein standard mix using Agilent Bio SEC-3 col-umns with (a) 300-, (b) 150-, and (c) 100-Å pore sizes. Eluent: 50 mM sodium phos-phate, 150 mM sodium chloride (pH 6.8). Peaks: 1 = thryroglobulin aggregates, 2 = thryroglobulin, 3 = IgA, 4 = globulins, 5 = ovalbumin, 6 = myoglobin, 7 = vitamin B12.


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and pH because these can influence the rate of formation and amount of aggregates. As part of the stability trials, formulations are tested at the end of their shelf life and this includes aggregate analysis.

Quality Control — Drug Substances and Drug ProductThe complex protein therapeutic must undergo multiple testing processes to ensure that it meets the defined quality requirements of product safety and efficacy. This includes testing for conformational and physical instabilities such as denatur-ation, unfolding, and aggregation because

these have implications for efficacy and immunogenicity (4,5). The methods used for the release of drug substances (clinical batches) during product development and drug product (production batches) after commercialization must be robust, provide the required sample throughput, and be validated according to guidelines from the International Conference on Harmonisa-tion (ICH) (ICH Q2) (6).

It is clear from the above that the pro-cesses required for manufacturing a bio-therapeutic are complex and that aggrega-tion could potentially change (increase or decrease) throughout the process.

Analytical Methods

Aggregates can be classified as soluble or insoluble, and the aggregation process can be reversible (noncovalent) or irreversible (covalent). Therefore, the analytical chal-lenges are immense with no single tech-nique capable of providing data across the range of solubilities and sizes, nanometer to millimeter (7).

SEC is used for the analysis and quan-titation of soluble aggregates that are non-covalent and irreversible in nature. This technique is a form of LC in which the separation is based on the hydrodynamic size of the protein in solution. SEC is suit-able for the analysis of aggregates in the size range of 1–50 nm and is considered robust and accurate when the method induces no changes to the aggregation state, and non-specific interactions with the column pack-ing media are eliminated.

The instability of the aggregate to changes in its environment or physical dis-ruption places exacting requirements on the method of analysis if accurate quantitation of aggregate content is the desired outcome. Just as with the manufacturing process, the method of analysis can also influence aggregation. Consideration must be given to the temperature, pH, and ionic strength of the eluent as well as sample preparation, concentration, agitation, and filtration if the data generated is to be representative of the

“sample.” In the following sections, choices for the development of SEC methods are explored and examples are presented to illustrate some of the pitfalls and consider-ation for development of an SEC method that matches the data need.

Size-Exclusion Chromatography

The Separation Mechanism In SEC, the molecules are separated based on differences in size in solution — with the elution order from largest to smallest. Figure 1 shows a schematic of the differential flow paths as a function of molecular size and the associated chromatogram. To achieve suf-ficient differentiation in residence time in the column, the SEC media must be porous and have a high pore volume.

The pore size of an SEC column will define the molecular sizes that can be resolved — anything that is bigger than the pores will be excluded and will be eluted at the exclusion volume of the column, and the smallest molecule that does not interact with the packing will be eluted at the total

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Figure 4: The effect of �ow rate on the resolution of a monoclonal antibody and its di-mer. Column: 150 mm × 7.8 mm, 3-µm Agilent Bio SEC-3 300 Å; eluent: 150 mM sodium phosphate, 100 mM sodium sulfate; �ow rates: 1.0, 1.5, and 2.0 mL/min.


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permeation volume — these two points define the elution volume and resolving range of the column. In the example shown in Figure 2 of the separation of the BioRad protein test mix, the 300-Å pore size col-umn resolves all seven components in the mix. With the smaller pore sizes, 150 Å and 100 Å, the larger sample components are eluted together at the exclusion point — with the 150-Å pore size column the thyro-globulin aggregates, thyroglobulin, and IgA are excluded and eluted together, and when the pore size is reduced further to 100 Å the globulins are also excluded. Therefore, the correct pore size must be selected based on the size of the proteins being resolved and the information required. To increase reso-lution, SEC columns are run in series, either combining the same pore size or different pore sizes to increase the resolving range.

Speed of Analysis When screening multiple samples, such as when looking at cell culture optimization or process variables on the level of aggrega-tion, the speed of analysis becomes critical. This is one of the drivers for the use of small particles, such as sub-2-µm particles, for small-molecule analysis. There is also inter-est in the use of sub-2-µm particles for the analysis of biomolecules, but there is one

significant difference that can impact the effectiveness of this strategy — the size of the molecule. When reducing the particle size, the porosity of the frit used to retain the media also decreases, nominally 0.5 µm, and the interstitial space in the packed col-umn decreases. Figure 3 compares the inter-stitial space for 3 µm and 1.7 µm particles. There is a large range in protein hydrody-namic radii, but typically those monomeric proteins of biopharmaceutical importance have hydrodynamic radii of between 2 nm and 13 nm (8); the aggregates are consider-ably bigger. With SEC, where the primary application is the analysis and quantitation of monomer, dimer, and multimers of pro-teins, there is a risk of shear forces causing changes to the sample composition or the column acting as a filter and trapping the multimer. In both of these examples, the relative composition of the samples will be altered by the SEC column.

An alternative way to reduce the analysis time is to use a shorter column and increase the flow rate. However, with SEC, where the separation between two molecules depends on the difference in the residence times, reducing the column length will impact resolution and, in practice, very short col-umns cannot be used. The separation of a monoclonal antibody obtained using a 150

mm long SEC column is shown in Figure 4. The 150-mm column is capable of resolv-ing the monomer from the dimer, but as the flow rate is increased from 1.0 mL/min to 2.0 mL/min the resolution decreases from 1.53 to 1.13; the percentage of dimer cal-culated was constant at 0.64%. As the size of the molecule increases, its mass transfer decreases with the effect that the peak width increases and impacts the resolution, and monomer efficiency decreases from 3510 to 1917 plates at 2 mL/min. This approach is suitable when high sample throughput is required (such as in the screening of cell cultures and the optimization of process conditions) because the time required for analysis is reduced to under 4 min. However, increasing the flow rate is not universally possible as some proteins, such as the clot-ting factors, aggregate in liquid streams at high flow rates and block the column. With the vast range of proteins it is important to understand the physicochemical stability and function requirements to ensure that the analytical method is fit for purpose and does not result in “changes” to the sample.

Quantitation of Soluble Aggregates, Dimers, and OligomersThere is no regulatory defined limit for the level of soluble aggregates in a biopharma-ceutical preparation because the acceptable level relates to its physicochemical char-acteristics, efficacy, and immunotoxicity. Therefore, the acceptable amount must be determined for each protein and quality control (QC) testing must be sufficiently robust and sensitive to quantify the level of aggregation at this limit.

Sample PreparationAs mentioned previously, protein aggrega-tion is influenced by environmental condi-tions and, therefore, if the requirement is to quantify the level of monomer, dimer, and higher aggregates then attention must be given to how the sample is prepared, so that the level of aggregation is not altered by the sample preparation method. Table I sum-marizes the SEC quantitation of a monoclo-nal antibody, as well as its dimer and higher aggregates. It also shows the effect of vari-ous sample treatments on the quantitation. With the monoclonal used in this study, sonication of the sample increased the monomer content and reduced the amount of both the dimer and the higher aggre-gates. The same observation was made for

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Figure 5: Size-exclusion analysis of bovine IgG with UV absorbance (280 nm, orange trace), light-scattering (90°, red trace), and refractive-index (blue trace) detection. Col-umn: 300 mm × 7.8 mm Bio SEC-5 500 Å; instrument: Agilent 1260 In�nity Bio-inert Quaternary LC with Agilent 1260 In�nity GPC/SEC Multi Detector Suite; mobile phase: 50 mM sodium phosphate, 250 mM sodium chloride (pH 7.0); �ow rate: 1.0 mL/min.


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the sample temperature, indicating that the dimer and the higher aggregates were dis-sociated at elevated temperatures to give a higher concentration of monomer.

Accuracy and PrecisionThe aggregate levels above which adverse reactions may be encountered can be a frac-tion of 1% in a biopharmaceutical prepara-tion. The analytical methods used for the analysis must, therefore, have a sufficiently low limit of quantitation (LOQ) and limit of detection (LOD) to provide accuracy and precision at these limits (9). As part of the development of an SEC method, robustness must be determined. Experi-ments must be carried out to demonstrate that the separation is based on size in solu-tion, there are no nonspecific interactions, and the analysis conditions, including those used to prepare the sample, do not change the amount or type of aggregates present in the sample. This can be done as part of a method robustness study (9).

Multiple Detectors When dimer and higher aggregate levels are very low, the LOQ and LOD achieved using UV or diode-array detection may not be sufficient. When light scattering is combined with SEC there is an increase in sensitivity for the dimer and higher aggre-gates. The added benefit of using an on-line scattering detector is that when used with a concentration detector such as a refractive-index detector, absolute molecular weights are obtained for homogenous peaks and the molecular weight profile is obtained where multiple species are present in a single peak. Figure 5 shows the elution profile of a bovine

IgG using refractive-index, UV (280 nm), and light-scattering (90°) detection. Figure 5 clearly shows the change in relative peak heights of the monomer, dimer, and higher aggregates when light-scattering detection is used compared to UV at 280 nm. A column with a 250–300 Å pore size would typically be used for the analysis of this size of protein when UV detection was used, but in this case a 500-Å pore size was needed to resolve the low level of higher aggregates that are now visible because of the increased sensi-tivity of the light scattering detector for the larger protein aggregates.

Conclusions

Aggregation of biotherapeutics is undesir-able because it can impact the process eco-nomics and efficacy of the product and can also cause adverse immune responses. Dur-ing manufacturing process development much emphasis is put on understanding the mechanism of aggregation for the target protein and on finding ways to control and reduce the folding and aggregation of the protein. SEC is widely used to study aggre-gation and characterize protein aggregates and is also suitable for quantifying levels of soluble aggregates in a preparation. It has been routinely adopted as one of the meth-ods of choice for the characterization of biopharmaceuticals and is applied through the process development stages and as one of the release criteria tests in QC. The tech-nique can be run to achieve fast separations by using short columns at higher flow rates or by using smaller particle sizes and for increased resolution multiple columns can be run in series and accurate quantitation can be achieved. By using a combination

of SEC and light-scattering detection, the sensitivity for large aggregates is improved, thereby enabling a reduction in the detec-tion levels and obtaining information about the molecular weight of the aggregations.

References

(1) J. Engelsman, P. Garidel, R. Smulders, H. Koll, B. Smith, S. Bassarab, A. Seidl, O. Hainzl, and W. Jiskoot, Pharm. Res. 28, 920–933 (2011).

(2) J. Patel, R. Kothari, R. Tunga, N.M. Rit-ter, and B.S. Tunga, BioProcess Int. 1, 20–31 (2011).

(3) W. Wang, Int. J. Pharmaceut. 185, 129–188 (1999).

(4) S. Hermeling, D.J.A. Crommelin, H. Schellek-ens, and W. Jiskoot. Pharm. Res. 21, 897–903 (2004).

(5) M.E.M Cromwell, E. Hilario, and F. Jacobson, The AAPS Journal 8(3), article 66, E572–E597 (2006).

(6) International Conference on Harmonisation, ICH Q2(R1), Validation of Analytical Proce-

dures: Text and Methodology (ICH, Geneva, Switzerland, 1994).

(7) T. Arakawa, J.S. Philo, D. Ejima, K. Tsumoto, and F. Arisaka, BioProcess Int. 4(10), 32–42 (2006).

(8) J.K. Armstrong, R.B. Wenby, H.J. Meiseman, and T.C. Fisher, Biophys. J. 87(6), 4259–4279 (2004).

(9) M.S. Palaniswamy, Agilent Application Solu-tion, Publication No. 5991-0835EN (2012).

Linda Lloydis currently the BioColumns Product Manager at Agilent Technologies with responsibility for global strategy. She has more than 30 years experience in GPC/SEC and HPLC and is the author of more than 40 scientific papers and conference presentations. Experience in biomolecule analysis and purification was gained at Polymer Laboratories and The University of Birmingham where she was involved in the design and development of new materials for biomolecule separations and the development of methods, including HPLC, to characterize bioactive proteins and polysaccharides. Direct correspondence to: [email protected] �



Table I: Quantitation of a monoclonal antibody, monomer, dimer, and high-er aggregates with different methods of sample preparation

Monomer (%) Dimer (%) Higher Aggregates (%)

Sample Sonication

Before sonication 76.1 16.5 7.4

After sonication 83.8 12.3 4.0

Sample Temperature

7 °C 76.1 16.5 7.4

15 °C 78.2 15.8 6.0

25 °C 80.3 14.4 5.3

35 °C 83.6 12.4 4.1

Sample Concentration

1 mg/mL 77.1 16.1 6.8

2 mg/mL 76.5 16.8 6.7

4 mg/mL 76.0 16.8 7.2


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Joseph L. Glajch, Momenta Pharmaceuticals. Direct correspondence to: [email protected]

Impact of New Columns on Drug Development

This article will examine the development of new types of

columns based on different particle types, sizes, and other

physical characteristics and how they can improve the speed

and efficiency of high performance liquid chromatography

used to support more expansive and complicated analyses.

Liquid chromatography (LC) has always been an important ana-lytical technique in the drug

development process. For small-mol-ecule drugs, the use of high perfor-mance liquid chromatography (HPLC) for assay and impurity analyses, anal-ysis of process samples, support of toxicology and clinical studies, and stability monitoring methods plays a crucial role in the entire development. Many of the separations for this class of drugs are based on reversed-phase HPLC and sometimes other tech-niques, such as chiral analysis. For biological drugs, the use of HPLC is even more important and expands to the use of size-exclusion chromatog-raphy, ion-exchange chromatography, and other specific techniques such as affinity chromatography.

The recent Affordable Health Care Act of 2010 in the United States provided a pathway for the develop-ment of biosimilar products, similar to the generic drug pathway for tra-ditional drugs that was established by the Hatch-Waxman Act of 1984. Although the specif ic requirements for approval of a biosimilar are still under development by the United States Food and Drug Administration (US FDA), it is clear from recent guid-ance and discussions that an increased emphasis will be placed on the ana-

lytical characterization as a part of the pathway for approval for biosimilars. A recent presentation by Kozlowski (1) highlighted the desire for an increased level of analytical and physical char-acterization in the development of biosimilar products.

This trend is creating an even greater need for high-eff iciency and high-performance analytics, many of which are based in part on HPLC. The development of new types of columns based on different particle types, sizes, and other physical char-acteristics will be a key contributor to the expanded use of HPLC to support these additional analytics. This arti-cle examines a few of these new tech-nologies and how they can improve the speed and eff iciency of HPLC used to support more expansive and complicated analyses.

Monolith Columns

One of the first needs for rapid HPLC analyses is the support of process development for biological molecules. Many large proteins are produced in cell-based fermentation systems. These cell culture production systems require signif icant process develop-ment to optimize the input starting materials, feed rates, and frequencies, and other parameters to generate protein at a commercially viable concentration


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in the final product. The need to con-tinuously and rapidly monitor protein concentration (often referred to as titer) is important to initially aid in the selection of a proper clone to be

used and then as additional process development work continues.

Silica-based monolith HPLC col-umns were developed in the early 1990s (2) as an alternative to tradi-

tional porous HPLC columns. These columns use channels rather than pores for f low through the column and are less likely to clog with other materials in the sample, such as cell debris in a cell culture sample. In addition, they rely on convective mass transfer leading to f low rate–independent separations that can provide very rapid analysis of high-molecular-weight samples such as proteins. A specif ic example of this is a monolith column, which contains protein A bound to the surface of the monoliths. Protein A selectively reta ins IgG proteins in an a f f in-ity type separation, therefore, these columns can be used to separate the IgG (the protein of interest in the cell culture) from other proteins and cell debris made during the cell culture process.

An example of this type of analy-sis is shown in Figure 1. In this case, the blue trace is a 0.5 mg/mL sample of the originator drug (traztuzumab) injected as the drug product. The red

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trace is a cell culture sample from one clone that was centrifuged for 5 min at 5000 g and the supernatant was directly injected onto an Agilent Biomonolith Protein A column (5.2 × 4.95 mm). The IgG protein is retained on the column while the other cell proteins and debris are eluted rapidly in less than 20 s in the initial mobile phase. A change to the mobile-phase B (citric acid buffer) elutes the IgG protein with a total analysis time of less than 2 min. Because multiple clone samples are often analyzed at once during clonal screening, this rapid analysis permits the screening of dozens or hundreds of samples quickly.

Small-Particle (Sub-2-µm)

Columns for High Efficiency

and Speed

A major development in column tech-nology has been the introduction of small (sub-2-µm) particles and the related equipment to handle the high pressure drop of these columns. Compared to classical 5-µm packed columns, these columns have per-mitted separations with much higher resolution, faster analysis times, or both, and have been widely applied in support of drug development. An

example of this speed is the analysis of the N-glycan profile of monoclonal antibodies (mAbs). Most monoclonal antibodies are glycosylated at one or more sites on the protein backbone. The glycosylation of one or more asparagine (N) residues is a com-mon post-translational modification (PTM) that must be measured. This is necessary either to develop a new biological drug or develop a biosimi-lar that closely matches the glycosyl-ation pattern of the innovator drug. Because the glycosylation is a PTM in the cell culture process, it can vary from batch-to-batch even in the same process. This variation creates the need to analyze multiple samples over many lots to establish the pattern or range of glycosylation; hence the need for an efficient and rapid way to assess this PTM.

Rapid N-Glycan Analysis

N-Glycan analysis is done by f irst treating the glycosylated protein with the enzyme N-glycanase to cleave the glycans from the protein backbone. Because glycans do not contain a good chromophore for UV detection, label-ing with a tag such as the f luorescent label 2-aminobenzamide is performed, followed by HPLC separation and

detection of the f luorescently tagged glycans.

Figure 2 shows the major glycan species that are found in many IgG proteins. Figure 3 shows the separa-tions of the N-glycans released from samples of rituximab and bevaci-zumab. Separat ion and quantita-tion of these species is done in this example using a Waters Acquity BEH Glycan column (100 mm × 2.1 mm; 1.7-µm particles) operated in hydro-phil ic-interaction chromatography (HILIC) mode. The separation is performed with a column tempera-ture of 60 °C and a f low rate of 1.0 mL/min using an acetonitrile–100 mM ammonium formate (pH 4.4) gradient with a total elution time of 10 min (for full experimental details see reference 4). Traditionally, these separations were done using more standard 5-µm columns and required 30–60 min per analysis. Using the small-particle and ultrahigh-pressure l iquid chromatography (UHPLC) systems, the complete analysis can be accomplished in less than 10 min, which is ideal for the analysis of mul-tiple lots or multiple process samples often required.

Rapid Amino Acid Analysis

Another need to support process devel-opment is amino acid analysis done during process optimization. This can be done to analyze the final protein product, precursors, or even the feed solution used in the process for spe-cific amino acids. Analysis of multiple samples is often required with very short turnaround to support process development and optimization.

Amino acid analysis of proteins has been done routinely since f irst reported by Moore and Stein in 1951 (5). Hydrolysis of the protein to free amino acid followed by HPLC using either pre- or post-column derivatiza-tion has routinely been done, with a typical separation time of 40–50 min per sample. An example of this sepa-ration is shown in Figure 4a using a standard HPLC column and system to provide baseline separation of all the amino acids. The use of a sub-2-µm column with a UHPLC system using similar separation conditions gives the

1

GO

9

A1F

Fucose

Galactose

Mannose

N-acetylgucosamine

N-acetylneuraminic acid 10

A2F

11

Man5

12

Man6

2

GO-N

3

GOF

5

G1

6

G1F

7

G2

8

G2F

4

GOF-N

Figure 2: N-Glycan structures, peak ID numbers, and notations. The peak ID numbers identify the associated structure on the N-glycan pro�les in Figure 3. Note that on the pro�le 6 and 6’ are G1F[6] and G1F[3], respectively. Adapted from reference 3.


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same separation, shown in Figure 4b, in only 10 min.

The overall analysis of a single sam-ple still requires multiple hours (pri-marily because of the sample digestion and other sample preparation time). However, because multiple samples can be digested in parallel, the sav-ings of HPLC analysis time can mean a signif icant overall time savings if multiple samples are required at once.

Superficially Porous

Particles for High Efficiency

and Rapid Separations

The use of small-particle columns (<2 µm) as described above can improve both the speed and efficiency of HPLC; however, another approach is to use superficially porous (sometimes called pellicular) particles of 2.7–4 µm that have a solid core and a “shell” of 0.4–0.8 µm outside the solid core. These

materials combine the high efficiency of the <2-µm particles (because of the effective separation of the 0.6-µm shell and their narrow particle size distribu-tion) with lower back pressures of 3–4 µm particles. The pellicular particles were first introduced in the late 1960s with 30–40 µm size and thin (2-µm) shells, but they fell out of favor after totally porous materials in the 5–10 µm size were produced. Superficially porous particles of 5–6 µm were rein-troduced in 2000 (6) for proteins and peptides with smaller cores and shells to extend this technology to smaller particles and, more recently, they were extended to the 2.7–4 µm range. A few examples of the use of this technology are described here to illustrate their use in drug development and process work.

Tryptic Peptide Analysis

Another major need in the develop-ment of biological drugs and biosimi-lars is in the separation of tryptic peptide maps of the intact protein. As an example, an IgG can be deglycosyl-ated and then digested with trypsin to generate a tryptic map, which can be a fingerprint of the protein and is often used as an identification test from lot to lot. A digest of a typical monoclonal antibody of MW 150 kDa produces approximately 60 tryptic peptides (depending on sequence), which can be separated by reversed-phase HPLC gradient elution using conventional HPLC columns. However, some of the amino acids contain PTMs such as oxidation of methionine, glycation of lysine, and deamidation of asparagine. These PTMs result in more peptides at lower levels that need to be separated from each other and the major species to identify and quantitate these lev-els. The use of higher-efficiency col-umns using smaller particles permits an expansion of the peak capacity of a typical tryptic map and the separation of many of these additional peptides.

An example of the use of these materials is shown in the top half of Figure 5 for the trypsin peptide map of a monoclonal antibody. The top chromatogram uses a 150 mm × 2.1 mm Advance Bio Peptide Map-ping column (Agilent Technologies)

8.00

(a)

(b)

1.50 2.00

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Q Asp Se

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3 Arg

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Gly

Asp G

lu

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r

Ala P

ro

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Lys

Tyr

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Leu

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e

Ph

e

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Time (min)

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10.00 12.00 14.00 16.00 18.00 20.00 22.00 24.00 26.00 28.00 30.00 32.00 34.00 36.00 38.00

0 2.2 2.4 2.6 2.8

4 1 11 & 5 7 12

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9 10 2 4 1 11 & 5

6

33

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Re

spo

nse

(R

FU)

Re

spo

nse

(R

FU)

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4.6 4.8 5.0 5.2 0 2.4 2.6 2.8 3.0 3.2 3.4 3.6 3.8 4.0 4.2 4.4 4.6 4.8 5.0 5.25.4 5.6 5.8 6.0 6.2 6.4 6.6 6.8

Figure 4: Comparison of amino acid separation on two different columns and sys-tems. (a) 150 mm × 3.9 mm AccQ-Tag column (Waters) (4-µm particle size) with a flow rate of 1.0 mL/min and a 50-min cycle time. (b) 100 mm × 2.1 mm AccQ-Tag Ultra column (1.7-µm particle size) with a flow rate of 0.7 mL/min and a 10-min cycle time. Both separations used gradient elution with proprietary eluents (AccQ-Tag and AccQ-Tag Ultra); total gradient times for the separations are 45 min and 9.5 min, respectively

Figure 3: Representative N-glycan pro�le of (a) rituximab and (b) bevacizumab. Peak numbers correspond to N-glycan structures in Figure 2. Adapted from reference 3.


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containing 2.7-µm particles with a 0.5-µm shell. The 75-min gradient run is similar to that obtained using a more traditional 3- or 5-µm totally porous particle, but achieves a higher resolution separation (in this case 59 major peaks observed in the 70-min run) with a moderate back pressure of 211 bar. A similar separation could be achieved with a totally porous 1.7-µm column of similar chemistry; how-

ever, the back pressure of the column would be substantially higher even at a f low rate of 0.2 mL/min.

If this comparison were the only difference, either a totally porous 1.7-µm column or a porous shell 2.7-µm column could be used to perform this separation with equal efficiencies and time of analysis. However, the porous shell offers one additional advantage not often used by most methods related

to an optimal gradient separation. The initial conditions for the 150 mm � 2.1 mm column were developed by scaling the f low rate from a 1.0 mL/min from a 150 mm � 4.6 mm column down to 0.2 mL/min for a 150 mm � 2.1 mm column (scaled down by a factor of five to account for the smaller inner diameter). Although this is an appro-priate scale factor for isocratic separa-tions, gradient elution separations are more complex and require scaling both f low rate and gradient characteristics to match the separation conditions changing from column to column. The important factor is to keep the appar-ent k value (often referred to as k*) con-stant from one condition to another. According to equation 1 (7), k* can be calculated as:

k* = 0.87 � tG � F/�Ø � Vm � S [1]

where tG is the gradient time; F is the column f low rate; �Ø is the gradient % change; Vm is the column volume, and S is a parameter related to solute molecular weight. For the purposes of this example, S can be assumed to be approximately 10 for the peptides of interest.

An original “traditional” separation using a 150 mm � 4.6 mm column with 3–5 µm particles would have tG of 75 min; F = 1.0 mL/min; �Ø of 0.3 (10–40%); Vm of 1.5 mL, so k* would be 14.5. Since a k* value of 2–10 is optimal, the traditional separation would have been reasonable for overall resolution.

If the translation to a smaller bore column (150 mm � 2.1 mm) only included the change in f low rate to 0.2 mL/min, k* would be calculated by equation 2 (very similar to that for the traditional column)

k* = 75 � (0.2/0.3) � 0.33 � 10 = 13.2 [2]

Since k* is the important parameter for a good gradient separation and a smaller k* could provide an equivalent or better separation, the porous shell column offers an additional advantage of increasing the f low rate F (from 0.2 to 0.6 mL/min) with a concomitant decrease in the gradient time (from 75 min to 14 min) and a small decrease in

160

Ab

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an

ce (

mA

U)

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mA

U)

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120

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80

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0 2 4 6 8 10 12Time (min)

Time (min)20 30 40 50 60

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%B

50%59 peaks, 211 bar

57 peaks, 433 bar

%B

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20

0

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6.0

5.0

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2

HC

4

HC

2

HC

1

HC

5HC

3

LC1

5.0

LC = light chain

HC = heavy chain

7.5 10.0 12.5 15.0 17.5

Time (min)

Ab

sorb

an

ce (

mA

U a

t 280 n

m)

Figure 5: Comparison of separation of tryptic peptides using different gradient and �ow conditions. (a) 150 mm � 2.1 mm AdvanceBio Peptide Mapping column; �ow rate: 0.2 mL/min; gradient time: 14 min. (b) 100 mm � 2.1 mm AdvanceBio Peptide Mapping column; �ow rate: 0.6 mL/min; gradient time: 75 min. Mobile-phase A: 0.1% formic acid in water; mobile-phase B: 90% acetonitrile in water with 0.1% formic acid; gradient: 3–35% B. (Courtesy of Agilent Technologies.)

Figure 6: Liquid chromatography–mass spectrometry (LC–MS) separation of mAb IgG1 light and heavy chains. Column: 100 mm � 2.1 mm Halo Protein C4; gradient: 29–32% B in 20 min; mobile-phase A: 0.5% (v/v) formic acid with 20 mM ammonium formate; mobile-phase B: 45% acetonitrile, 45% isopropanol–0.5% (v/v) formic acid with 20 mM ammonium formate; temperature: 80 °C; �ow rate: 0.4 mL/min; instru-ment: Shimadzu LCMS 2020; detection: 280 nm. (Courtesy of Advanced Material Technology.)


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column length (to 100 mm) resulting in a k* calculated in equation 3,

k* = 14 × (0.6/0.3) × 0.22 × 10 = 11.1 [3]

Thus, maintaining a reasonable k* value (and separation efficiency) while decreasing the overall run time. The comparison chromatogram in the bot-tom part of Figure 5 shows the result

— an almost identical separation of 57 peaks with a markedly reduced run time of 14 min. Since the column back pressure is still 433 bar, this can be accomplished on either standard HPLC or UHPLC equipment.

Large-Pore Superficially Porous

Particles for Protein Separations

Many of the superficially porous par-ticles were designed with pore sizes in the range of 80–160 Å (8–16 nm) with a shell thickness of 0.5–0.6 µm (500–600 nm). To separate even larger materials such as proteins, columns with pore sizes of �300 Å are needed. In addition, a thin shell is desirable to create a short diffusion path for these larger molecules and maintain high efficiency of separation.

Schuster and colleagues (8) described the development and synthesis of such materials and concluded that 3.4-µm particles with 400-Å pores and a 0.2-µm shell thickness provides a good compromise of pore size, shell thick-ness, and mass loadability to provide improved separations. A good example for the use of such materials is the analysis and quantitation of the light chain and heavy chain portions of an IgG monoclonal antibody, including the separation of different known vari-

ants because of different glycosylation patterns on the heavy chain and other PTMs such as oxidation or pyro-glu-tamic acid modification.

A typical procedure for this analy-sis involves denaturation of the intact protein, reduction, and alkylation to produce light chain, heavy chain, and any variants because of post-transla-tional modification. Separation is nor-mally done only into light chain and heavy chain regions, which are then further analyzed by mass spectrom-etry (MS) to deconvolute the peaks into different mass variants and rela-tive abundances.

An improved separation of the indi-vidual light chain and heavy chain variants would provide a better under-standing of the actual distribution of these species. Figure 6 shows the sepa-ration of light chain and heavy chain variants along with the mass spectral data for the individual peaks. The improved resolution, especially of the light chain variants, permits a more robust comparison of these in mul-tiple samples from different lots and sources.

The peaks can also be analyzed by MS as the individual species and the component masses can be deconvo-luted using the appropriate software. Table I shows the results for the three light chain peaks and the five heavy chain peaks in this example.

Conclusions

The use of columns of different types, particle sizes, and pore sizes can pro-vide signif icant advantages in both speed and resolution for the analysis of complex samples during drug discov-ery and development. Given the com-plex nature of many biological drugs and the need to more fully characterize these for both initial development and biosimilar development, the increased use of these types of columns and sys-tems will be crucial to these efforts.

Acknowledgments

I would like to thank Maureen Joseph and James Martosella of Agilent, Jus-tin Hyche and Ted Haxo of ProZyme, Bill Warren and Priya Jayaraman of Waters, and Stephanie Schuster and Jack Kirkland of Advanced Materials

Technology for their help in obtaining examples used in this paper.

References

(1) S. Kozlowski, “Biosimilar Biological Prod-ucts: Overview of Approval Pathway under the Biologics Price Competition and Inno-vation Act of 2009,” presentation at the 17th Symposium on the Interface of Regu-latory and Analytical Sciences for Biotech-nology Health Products (WCBP 2013).

(2) K. Nakanishi and N. Soga, J. Am. Ceram.

Soc. 74, 2518–2530 (1991).(3) S. Fuller, T. Haxo, J. Hyche, M. Kimzey,

S. Lockart, S. Pourkaveh, Z. Szabo, J. Truong, J. Wegstein, and V. Woolworth,

“Assessing the Variability of an Innovator Molecule N-Glycan Profile,” poster at the 18th Symposium on the Interface of Regu-latory and Analytical Sciences for Biotech-nology Health Products (WCBP 2014).

(4) www.prozyme.com/protocols/.(5) S. Moore and W.H. Stein, J. Biol. Chem.

192, 666 (1951).(6) J.J. Kirkland, F.A. Truszkowski, C.H.

Dilks Jr., and G.S. Engel, J. Chromatogr.

A 890(1), 3–13 (2000).(7) L.R. Snyder, J.J. Kirk land, and J.W.

Dolan, Introduction to Modern Liquid

Chromatography, 3rd ed. (Wiley, Hobo-ken, New Jersey, 2010), p. 412.

(8) S.A. Schuster, B.M. Wagner, B.E. Boyes, and J.J. Kirkland, J. Chromatogr. A 1315,

118–126 (2013).



Joseph L. Glajchis Director of Analytical Development at Momenta Pharmaceuticals. He received his AB in Chemistry at Cornell University and PhD in Analytical Chemistry at the University of Georgia under L.B. (Buck) Rogers. He has held technical and R&D management positions at DuPont, Bristol-Myers Squibb, and Momenta with emphasis on HPLC column and method development and pharmaceutical development and analysis. He is also a member of the Editorial Board of LCGC North America. Direct correspondence to: [email protected] �

Table I: Deconvoluted mass values for each peak in Figure 6

Peak Mass (Da)

LC1 23,204

LC2 23,192

LC3 23,303

HC1 50,539

HC2 50,424

HC3 50,668

HC4 50,680

HC5 28,862


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Mirlinda Biba*,†, Bing Mao*, Christopher J. Welch*, and Joe P. Foley†

*Department of Analytical Chemistry, Merck Research Laboratories, Rahway, New Jersey; †Department of Chemistry, Drexel University, Philadelphia, Pennsylvania. Direct correspondence to: [email protected]

Liquid Chromatography Methods for the Separation of Short RNA Oligonucleotides

Synthetic oligonucleotides have become increasingly popular as a

result of the recent discovery of ribonucleic acid interference (RNAi), a

natural process for silencing gene expression. As biomedical researchers

evaluate the use of antisense and small interfering RNAs (siRNAs)

as potential therapies for the treatment of disease, the need for

improved methods for the chromatographic separation and analysis of

oligonucleotides has become apparent. This article presents a review

of different liquid chromatography (LC) methods and strategies for

the chromatographic separation of short RNA oligonucleotides.

There has been considerable interest recently in the use of synthetic oligo-nucleotides as potential therapeutic

agents capable of suppressing the synthesis of specific proteins (1–3). Targeted “knock-down” of specific gene products using an antisense ribonucleic acid (RNA) strategy dates to the late 1990s (4). In this approach, a single-stranded oligonucleotide comple-mentary to the messenger RNA (mRNA) encoding a targeted protein leads to disrup-tion of ribosomal transcription and protein synthesis. In theory, antisense oligonucle-otides can be applied to any disease in which protein overexpression is detrimental, and a number of antisense oligonucleotides have been evaluated as potential therapies (5). The need for long complementary oli-gonucleotides and the stoichiometric nature of mRNA inactivation (1 antisense mole-cule:1 mRNA inactivation) places consider-able constraints on developing cost-effective antisense drugs.

The more recently discovered small interfering RNA (siRNA) mechanism for silencing gene expression involves a short double-stranded RNA molecule of about 21 base pair length, which activates the RNA interference (RNAi) silencing path-way (6,7), thereby achieving catalytic deg-radation of the target mRNA (one siRNA

molecule inactivates multiple mRNAs). An overview of the RNAi pathway for targeted gene silencing is illustrated in Figure 1.

The discovery of siRNA gene silencing in animals (8) and human cells (9) has led to a surge of interest in the use of siRNA for biomedical and drug development research. Many biomedical and pharmaceutical com-panies have become involved in the explora-tion of the preparation and use of siRNAs as potential therapies for the treatment of diseases such as cancer, macular degenera-tion, and viral infections (10).

Oligonucleotide and siRNA

Structure and Preparation

RNA is a biologically important molecule that consists of a long chain of nucleotide units. Each nucleotide contains a ribose sugar, a nitrogenous base, and a phosphate group. There are four bases in RNA: ade-nine (A), guanine (G), cytosine (C), and uracil (U) (Figure 2). Oligonucleotides are short, single-stranded RNA or deoxyri-bonucleic acid (DNA) molecules that can readily bind, in a sequence-specific manner, to their respective complementary oligonu-cleotides to form duplexes. Small interfer-ing RNA is a small double-stranded RNA (usually 21 nucleotides) with two nucleo-tide overhangs on each 3�-end. Each strand


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has a 5�-phosphate group and a 3�-hydroxyl group (Figure 3).

Various chemical modifications are often made to synthetic oligonucleotides to pre-vent attack by nucleases, which can lead to siRNA degradation and instability (11,12). Incorporation of either a fluoro or methoxy group into the 2� position of the sugar or the use of a phosphothioate linkage is commonly used to improve siRNA stabil-ity (Figure 2) (13). In the phosphothioate modification, oxygen in the phosphodies-ter linkage is replaced with a sulfur atom. This introduces an additional stereocenter into the molecule giving rise to two pos-sible diastereomers for every phosphothio-ate linkage, and making the resulting oligo-nucleotide sample mixtures highly complex and very difficult to chromatographically

resolve. All of these modifications help to improve oligonucleotide stability while retaining, and sometimes even increasing, their silencing activity. These modifications also tend to increase the hydrophobicity of the oligonucleotides, while also increasing the temperature at which the duplex melts (Tm) into its corresponding single strands.

Oligonucleotides are readily synthesized via stepwise synthesis using phosphorami-dite chemistry with automated solid-phase synthesizers (14). Although the individual synthetic process reactions can be very efficient and provide high yields, the total number of synthetic steps for making a 21-mer RNA can be more than 80 chemi-cal steps (with about four chemical steps for each cycle). Consequently, because of the accumulation of many small errors, the

final oligonucleotide product typically con-tains a variety of closely related impurities that can be very difficult to separate and remove during final product purification (15). Some of the most common impuri-ties include sequence deletions, such as n–1, n–2, and so on, where one or more nucleo-tide fails to attach to the sequence during synthesis. Additionally, depurination, oxi-dation, and other chemical modification or degradation of the nucleotide bases can lead to a variety of closely related impuri-ties that can be very challenging to resolve from the desired product. When dealing with double-stranded siRNAs, the sample mixtures can become even more complex with each strand introducing its own set of impurities. These impurities include mis-matched sequences and noncomplementary single stranded sequences. The presence of these impurities in a therapeutic mixture can lead to unwanted, nontargeted gene silencing, while the presence of any non-hybridized single-stranded RNAs can also lead to a decrease in therapeutic potency (40). Therefore, when developing siRNA therapeutics, one of the major challenges is ensuring good purity to minimize off-target gene silencing effects. Consequently, developing good chromatographic tech-niques is often critically important in the oligonucleotide drug development process.

Chromatographic

Separation of Oligonucleotides

When developing analytical methods for the separation of oligonucleotides, some of the unique features of these molecules need to be considered. First, the pKa of the phosphodiester linkage is 2, meaning that in aqueous solution above pH 4 these molecules contain one negative charge for every phosphodiester linkage — that is, a 21-mer oligonucleotide contains 21 nega-tive charges at pH 7. Therefore, traditional reversed-phase liquid chromatography (LC) conditions tend not to work well, and ion-pair reversed-phase LC or anion-exchange chromatography techniques need to be considered. Secondly, many single-stranded RNA oligonucleotides can form higher order (tertiary) structures including bends, loops, dimers, or other aggregates, and ele-vated chromatographic temperatures must be used to “melt” such structures, allowing for efficient analysis (16). Also, when ana-lyzing single-strands versus duplex samples, different methods must be considered, with

Cytoplasm

Long dsRNA

Dicer

siRNA

SyntheticsiRNA

Unwound passenger strand

mRNA cleavage

RISC assembly

Figure 1: RNA interference mechanism. Long dsRNA in the cytoplasm is cleaved into 21-mer strands (siRNA) by the protein, Dicer. Small interfering RNA is incorporated into the RNA-induced silencing complex (RISC), where the passenger strand is un-wound and degraded leaving the guide strand bound to RISC. The RISC-guide strand complex base-pairs with a complementary sequence of the mRNA and induces cleav-age of the mRNA, thereby preventing protein translation. Synthetic siRNAs can be introduced into the cell and achieve the same action in the RNAi mechanism.


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single-strand analysis typically taking place at temperatures >60 °C and duplex samples typically being analyzed at or below room temperature. A typical siRNA duplex LC analysis is shown in Figure 4. Finally, since RNA and DNA molecules both have a strong absorbance at 260 nm, these sam-ples are often analyzed using UV detection, although fluorescence and especially mass spectrometry (MS) detection (17) are also quite important for RNA analysis.

A number of chromatographic methods have been reported for the analysis and purification of oligonucleotides, includ-ing capillary gel electrophoresis (CGE) (18,19), anion-exchange high performance liquid chromatography (HPLC) (20–33),

ion-pair reversed-phase LC (34–51), and mixed-mode LC (52–56). These different LC techniques are reviewed in the follow-ing sections.

Ion-Exchange Liquid ChromatographyIon-exchange chromatography is an excellent method for separating charged molecules, and is a commonly used chro-matographic method for the separation of multiply charged oligonucleotides (20). In ion-exchange chromatography, separation is based on the differential electrostatic affinities of charged molecules for a charged stationary phase. For the separation of highly negatively charged oligonucleotides,

anion-exchange chromatography is used with positively charged stationary phases that exchange the negatively charged oli-gonucleotides through competition with anions from the mobile phase. As a result, longer oligonucleotides with greater charge are more strongly retained on the col-umn and shorter oligonucleotides, which have progressively fewer negative charges, have progressively shorter retention. This method is especially useful for the separa-tion of the very common N–x deletions of oligonucleotides (Figure 5a), but is less use-ful for detecting subtle changes on the full-length sequence that do not alter the total number of charges (Figure 5b).

Successful separation of small oligonucle-otides and their sequential analysis by anion-exchange HPLC was originally reported more than 30 years ago (21,22). In these studies, anion-exchange columns, such as Permaphase AEX (DuPont) or Partisil SAX (HiChrom), with a salt gradient using phos-phate or acetate buffers were used. Later, Pingoud and colleagues (23) showed the use of strong-anion-exchange LC for the excel-lent resolution of longer oligonucleotides (up to 64 bases) from their N–1 deletion products, using a Whatman SAX column and a phosphate salt gradient. Over the years, improved types of anion-exchange resins were developed to improve the resolu-tion of oligonucleotides with strong-anion-exchange LC. Alkylamine derivatized (24) and polyethyleneimine (PEI) coated (25,26) porous silica phases were prepared for the anion-exchange HPLC separation of oligo-nucleotides, where resolution of oligonucle-otides of up to 30 bases from their N–1 deletion products was obtained. Optimi-zation of the PEI bonding chemistry with quaternization of the ion-exchange matrix was shown to further increase the resolution of N–1 deletion products for up to 50-mer oligonucleotides (27).

For chromatographic separations using harsher conditions such as elevated col-umn temperatures and neutral-to-high pH mobile phases, the relative chemical insta-bility of silica-based stationary phases can be a significant disadvantage, because par-ticle erosion and column degradation can lead to a significant loss of performance over time (28). Consequently, PEI-coated porous zirconia stationary phases were pro-vided by Professor Carr for use as anion exchangers in the strong-anion-exchange LC of oligonucleotides (29). These stationary

RNA Bases

Purines

O

O

OO

O O

OOO–

O–

OO

OOO

O

O

OO

OO

5’

O

P

P

F

P

Base3

Base4

Base2

Base1

S

OH

OH

Phosphorothioate

H

NH2

NH2

H2N

N

NN

N

H

HH

H

Adenine (A)

Cytosine (C)

3’2’ -O-Me modi�cation

2’ -F modi�cation

Uracil (U)

Guanine (G)Pyrimidines

H

N

N

NN

N N

N

N

siRNA Structure

5’ PO4

5’ PO43’ OH

3’ OH

Figure 2: Oligonucleotide structure. Different modi�cations include phosphothioate backbone modi�cation where one oxygen atom on the phosphodiester backbone is replaced with a sulfur atom, and 2’-sugar modi�cations, such as 2’-F and 2’-O-Me. The four bases in RNA are adenine (A), guanine (G), cytosine (C), and uracil (U).

Figure 3: Small interfering RNA (siRNA) structure. A 21-mer siRNA is shown with two nucleotide overhangs on each 3’-end. Each strand has a 5’-phosphate group and a 3’-hydroxyl group. The siRNA duplex consists of two complementary strands, the sense (or passenger) and antisense (or guide) strands. In RNA, adenine base-pairs with uracil by forming two intermolecular hydrogen bonds (A–U) and guanine base-pairs with cytosine by forming three intermolecular hydrogen bonds (G–C).


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phases were able to provide single-nucleo-tide unit resolution for oligonucleotides of up to 50-nucleotide length. Furthermore, the zirconia-based stationary phases can be operated at elevated column temperatures of 75 °C, which allow for the highly advan-tageous elution of oligonucleotides with a low ionic strength mobile phase; those conditions lead to loss of performance and bed collapse with conventional silica-based columns.

The development of nonporous anion exchangers for use in strong-anion-exchange LC of proteins (30) and oli-gonucleotides (31) is another important milestone. These nonporous columns, such as TSKgel (Tosoh Biosciences), were pre-pared by introducing diethylaminoethyl (DEAE) groups into nonporous spherical hydrophilic resins with 2.5-µm diameter particles. The effects of some of the criti-cal chromatographic parameters in strong-anion-exchange LC for oligonucleotides were also studied. Examination of the elu-ent pH showed that pH should ideally be �8.5 for the separation of oligonucleotides. Furthermore, the range of pH 8.5–9.5 was shown to be better suited for separations based on differences in chain length, while a pH of 10.5 was better suited for separa-tion based on differences in base compo-sition. Because these stationary phases are chemically very stable, operating at high pH would not cause any problems. The addition of salts, such as sodium chloride or sodium perchlorate, also has an influence on chromatographic separation. Finally, other useful methacrylate-based (polymeric) anion-exchange stationary phases have also

been developed and successfully used for oligonucleotide separations (32–34).

Ion-Pair Reversed-Phase Liquid ChromatographyIon-pair reversed-phase LC is another very commonly used LC technique for the anal-ysis and separation of oligonucleotides (35). In ion-pair reversed-phase LC, negatively charged oligonucleotides interact with posi-tively charged alkylammonium ions in a way that permits the chromatographic sep-aration of the former on a reversed-phase stationary phase. Different mechanisms regarding the ion pair interaction have been proposed, depending on whether the ion-pair process occurs in the mobile phase (36) or in the stationary phase (37). In the first proposed mechanism, ion-pair formation occurs in the aqueous mobile phase, and the neutralized ion pair is then adsorbed onto the hydrophobic stationary phase, with retention controlled by the overall hydrophobicity of the ion pair. In the sec-ond mechanism, the unpaired ion (such as an alkylammonium ion) from the mobile phase is adsorbed onto the stationary phase, which then acts as an ion-exchange station-ary phase for the separation of charged oli-gonucleotides.

In ion-pair reversed-phase LC, in addi-tion to the charge–charge interactions, hydrophobic interactions from the indi-vidual bases also significantly contribute to the overall oligonucleotide retention. The hydrophobicity of oligonucleotide bases follows the order C < G < A < T (for DNA based oligonucleotides), with cytosine being the least hydrophobic base

(38,39). Therefore, in addition to the oli-gonucleotide length, the retention of oli-gonucleotides also depends on the specific base composition, where the overall hydro-phobicity is the sum of all the bases in the sequence. As such, ion-pair reversed-phase LC can be especially useful for separating impurities from changes in the full-length sequence such as depurinations and other chemical modifications on the bases. Sepa-ration of an oligonucleotide and a series of its deletions (N–1 to N–15) are shown in Figure 6a. This illustrates the retention of oligonucleotides by ion-pair reversed-phase LC in which the longer oligonucleotides are more retained, but overall retention is mostly governed by the base composition, where cytosine (the least hydrophobic) base deletions resulted in increased retention. Additionally, the ion-pair reversed-phase LC method provided some separation of very subtle differences on the oligonucle-otide sequence, such as the reversal of two neighboring bases at different locations along the sequence (Figure 6b).

Triethylammonium acetate (TEAA) is one of the most commonly used ion-pair reagents for the separation of oligonucle-otides because of its good separation effi-ciency (40,41). Typical TEAA concentra-tions in the aqueous solution are 100 mM at pH 7 with acetonitrile as the organic solvent. Gilar and colleagues (42) did an extensive study of the prediction of reten-tion for oligonucleotides with ion-pair reversed-phase LC using TEAA at pH 7 as the ion-pairing reagent. Using 39 dif-ferent oligonucleotides, they demonstrated the successful application of a model for the prediction of the mobile phase strength required to elute the oligonucleotides. Shal-low linear gradients of organic modifiers are typically used for these separations because studies have shown that there can be a sharp change in the retention factor (k) of oligonucleotides with a small change in the mobile phase strength. Gilar and colleagues (42) reported that the retention factor for a 15-mer oligonucleotide decreased from 100 to 13.5 to 3.2 with a very small change of mobile-phase composition from 8% to 9% to 10% acetonitrile.

Another very useful ion-pairing mobile phase is the combined hexafluoroisopro-panol (HFIP) and triethylamine (TEA) buffer system proposed by Apffel and col-leagues (43). This HFIP-based separation uses methanol as the organic modifier

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Figure 4: A typical UHPLC analysis of duplex 21-mer siRNA. Column: 100 mm × 2.1 mm, 1.7-µm dp Waters CSH C18; mobile-phase A: 400 mM HFIP–16.3 mM TEA in water (pH 7.9); mobile-phase B: methanol; segmented linear gradient: 25–33% B in 10 min, 33–36% B in 20 min, 36–60% B in 28 min, 3-min equilibration at 25% B; total run time: 31 min; �ow rate: 0.3 mL/min; detection: UV absorbance at 260 nm; column temperature: 15 °C.


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because HFIP is immiscible with aceto-nitrile and miscible with water, methanol, isopropanol, and hexane. Using 400 mM HFIP and adjusting the solution to pH 7.0 with TEA, comparable separations were obtained as those with the 100 mM TEAA mobile phases. The main difference, however, was in the electrospray ionization (ESI) performance for the MS detection of oligonucleotides. Comparison of different solvent systems, such as 400 mM HFIP adjusted to pH 7.0 with TEA, water, 100 mM TEAA (pH 7), and 25 mM TEA (pH 10), showed superior MS signal with the 400 mM HFIP buffer system, compared to significantly suppressed MS signal with the 100 mM TEAA buffer. This significant difference in MS detection was attributed to the dynamic adjustment of the pH in the ESI droplet as a function of the removal of the anionic counterion from the droplet

by evaporation. First, comparing the vola-tilities for the two different buffer systems, HFIP (boiling point [bp] = 57 °C) is more volatile than TEA (bp = 89 °C), with acetic acid being the least volatile (bp = 118 °C). Secondly, the weak acid and base system with HFIP and TEA maintains a stable pH at ~7.0. The pKa values of acetic acid, HFIP, and TEA are 4.75, ~9, and 11.01. Therefore, at pH 7.0, acetic acid is completely dissoci-ated and it cannot be removed by evapora-tion on the MS source, whereas HFIP is not charged and it can be evaporated freely. Furthermore, the mechanism proposed by Apffel suggests that during the separation, the TEA ions ion-pair with the negatively charged phosphates on the oligonucleotide backbone, because the more volatile HFIP is evaporated from the droplet surface caus-ing the pH on the surface to rise (~ pH 10). As the pH rises, the oligonucleotide–TEA

ion pair dissociates, and the oligonucle-otide can be desorbed into the gas phase. Additionally, the role of TEA is also very important in this mechanism because, in general, oligonucleotides have a high bind-ing affinity for Na+ and K+ cations on the polyanionic phosphate backbone and these cation adducts can diminish the sensitiv-ity for electrospray ionization. The use of a strong base such as TEA effectively sup-presses the sodium and potassium adducts formation by a displacement mechanism and consequently dramatically increases the ESI sensitivity. This HFIP and TEA mobile-phase system is now routinely used for oligonucleotide analysis with ion-pair reversed-phase LC and ESI-MS detection (43–50).

Gilar and colleagues (47) further evalu-ated the HFIP and TEA buffer system and showed that concentration of the ion-pair-ing TEA ion, rather than the concentration of HFIP, in the mobile phase plays a criti-cal role in the separation. They extensively studied the effect of the concentration of both, TEA and HFIP in the mobile phase by varying the TEA concentration from 0.56 to 31.4 mM range (pH 7–9), and the HFIP concentration from 12.5 to 400 mM, using oligonucleotides up to 30-mer length. They showed that the HFIP and TEA buf-fer system was most effective at 400 mM HFIP and 16.3 mM TEA concentration (where 16.3 mM TEA was the highest con-centration possible to dissolve in 400 mM HFIP at room temperature). Interestingly, they also showed that HFIP effectively dis-rupts any oligonucleotide secondary struc-tures and this buffer can be an efficient denaturant that allows for more efficient oligonucleotide separations.

As reported by Huber, Oefner, and Bonn (39,51–53), nonporous poly(styrene–divi-nylbenzene) (PS-DVB) particles with a diameter of 2.1 µm were prepared and successfully used for the ion-pair reversed-phase LC separation of oligonucleotides with methods using 100 mM TEAA as the ion-pairing reagent and a column tempera-ture of 50 °C. Other reversed-phase media such as porous C18 columns can also be used.

The mass transfer of relatively larger mol-ecules such as oligonucleotides is one of the major factors contributing to peak broad-ening on porous C18 stationary phases. A study using 50 mm × 4.6 mm columns packed with 5-, 3.5-, and 2.5-µm particles

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Isomer 5: 5’-GUCAUCACACUGAAUACCAUA-3’

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Isomer 3: 5’-GUCAUCACACGUAAUACCAAU-3’

Isomer 2: 5’-GUCACUACACUGAAUACCAAU-3’

Isomer 1: 5’-UGCAUCACACUGAAUACCAAU-3’

21-mer: 5’-GUCAUCACACUGAAUACCAAU-3’

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Figure 5: Chromatographic separation of oligonucleotide standards by strong-anion-exchange LC: (a) Separation of N–x deletion series, (b) separation of “base-�ip” iso-mer standards (25–29 min portion of chromatogram shown). Column: 150 mm × 4.6 mm, 3-µm dp Proteomix SAX-NP3 (Sepax Technologies); mobile-phase A: 80:20 (v/v) 10 mM Tris (pH 8)–ethanol; mobile-phase B: 600 mM sodium bromide salt concentra-tion in mobile-phase A; linear gradient: 20–80% B over 30 min with 5-min column equilibration at 20% B; �ow rate: 0.5 mL/min; column temperature: 60 °C; injection volume: 15 µL; detection: UV absorbance at 260 nm. Adapted with permission from reference 59.


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showed that the mass transfer in the sta-tionary phase had a major impact on the separation (42). To overcome this problem, columns containing smaller particles (�2.5 µm in diameter) and the use of ultrahigh-pressure liquid chromatography (UHPLC) instruments can be used to improve overall separation performance (40). Additionally, the use of core–shell C18 particle columns has been shown to significantly improve oligonucleotide separations compared to fully porous particles (54). Systematic eval-uation of different core–shell C18 columns showed the best separation with a sub-2-µm core–shell particle column. However, the long-term stability of these silica-based columns when operating at neutral pH and elevated column temperatures of >60 °C can be an issue.

Mixed-Mode Liquid ChromatographyBecause of differences in the separation mechanism between the strong-anion-

exchange LC and ion-pair reversed-phase LC methods, these approaches are some-what complementary and often both methods are used for complete analysis and characterization of oligonucleotide mixtures (55,56). As a result, mixed-mode chromatography columns possessing both reversed-phase and ion-exchange proper-ties have been prepared and evaluated for use in oligonucleotide separations (57,58). When using mixed-mode columns, oligo-nucleotides can experience ionic (such as in strong-anion-exchange LC) and hydropho-bic (such as in reversed-phase LC) interac-tions simultaneously. The dominating mode of interaction can be significantly influenced by the type of mobile phase used.

More recently, an evaluation of differ-ent Scherzo C18 mixed-mode (Imtakt USA) columns showed a significant ben-efit when using mixed-mode columns for separation of oligonucleotides (59). In addition to providing good separation of

typical oligonucleotide impurities, such as N–x deletions, which can be well sepa-rated by either strong-anion-exchange LC or ion-pair reversed-phase LC alone, these columns also showed an excellent separa-tion of very challenging isomeric oligo-nucleotides where one single nucleotide base was reversed with its neighboring base, affording separations that could not be achieved by either strong-anion-exchange LC or ion-pair reversed-phase LC alone. A mixed-mode chromatography separation of oligonucleotides with the Scherzo SM-C18 column and sodium chloride salt gradient is shown in Figure 7.

Future Approaches to the

Chromatographic Separation

of Oligonucleotides

As research on the biomedical uses of short RNA oligonucleotides continues, we can expect ongoing development of improved methods to chromatographically analyze and purify these fascinating compounds. Given the readiness with which these mol-ecules engage in Watson-Crick base pair-ing with complementary oligonucleotide strands, affinity chromatography–like approaches in which a complementary oli-gonucleotide stationary phase is used for the targeted retention and chromatographic separation of particular oligonucleotide products may be possible for the analysis, and especially purification, of RNA oli-gonucleotides. Affinity chromatography purification of oligonucleotide binding proteins has been carried with an oligo-nucleotide affinity column (oligo (dT)12-18 cellulose) (60) and more recently, with a stationary phase consisting of 2�-fluoro modified RNA covalently linked to aga-rose beads (61). Oligonucleotide hybridiza-tion has long been used in the formation of DNA microarrays and similar technologies (62,63), and can reliably be used for the selective capture of particular oligonucle-otide sequences. While these experiments involve only simple binding at lower tem-perature with release at elevated tempera-ture, true chromatographic separation may be possible on complementary oligonucle-otide stationary phases operated at elevated temperatures, or in the presence of mobile phase additives that make adsorption or desorption fast on the “HPLC time scale.” Such stationary phases could, in principle, be tailor-made for analytical or purification tasks such as the selective binding of desired

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N-15: 5’-ACCAAU-3’N-14: 5’-UACCAAU-3’N-13: 5’-AUACCAAU-3’N-12: 5’-AAUACCAAU-3’N-11: 5’-GAAUACCAAU-3’N-10: 5’-UGAAUACCAAU-3’N-9: 5’-CUGAAUACCAAU-3’N-8: 5’-ACUGAAUACCAAU-3’N-7: 5’-CACUGAAUACCAAU-3’N-6: 5’-ACACUGAAUACCAAU-3’N-5: 5’-CACACUGAAUACCAAU-3’N-4: 5’-UCACACUGAAUACCAAU-3’N-3: 5’-AUCACACUGAAUACCAAU-3’N-2: 5’-CAUCACACUGAAUACCAAU-3’N-1: 5’-UCAUCACACUGAAUACCAAU-3’21-mer: 5’-GUCAUCACACUGAAUACCAAU-3’







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Figure 6: Chromatographic separation of oligonucleotide standards by ion-pair re-versed-phase LC: (a) Separation of N–x deletion series, (b) separation of “base-�ip” isomer standards (11–15 min portion of chromatogram shown). Column: 150 mm × 4.6 mm, 3.5-µm dp XBridge C18 (Waters); mobile-phase A: 100 mM TEAA in water; mobile-phase B: 100 mM TEAA in acetonitrile; linear gradient: 5–10% mobile phase B over 15 min with 5-min column equilibration at 5%; �ow rate: 1.5 mL/min; column temperature: 65 °C; injection volume: 15 µL; detection: UV absorbance at 260 nm. Adapted with permission from reference 59.


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target oligonucleotides, or of an otherwise difficult-to-remove isomeric or closely related impurity. A preliminary investiga-tion showed that the use of RNA-based stationary phases for selective purification of short RNA sequences suffers from rapid degradation of the RNA stationary phase at elevated column temperatures (64), a problem that could potentially be solved by the use of more thermostable DNA-based stationary phases or even the use of com-plementary phases based on peptide nucleic acids (PNAs), which are significantly more thermostable than either RNA or DNA, but retain the ability to form complemen-tary duplexes (65).

Conclusions

Developing good chromatographic meth-ods for the accurate and sensitive analysis and separation of oligonucleotides is a criti-cal part in biomedical investigations involv-ing antisense and siRNA oligonucleotides. In this review, we have surveyed different

LC approaches including strong-anion-exchange, ion-pair reversed-phase, mixed-mode, and affinity liquid chromatography. Strong-anion-exchange LC is one of the most often used methods in which sepa-ration of oligonucleotides is mainly based on the different charges, making this tech-nique especially useful for the separation of the very common N–x deletion impu-rities. Ion-pair reversed-phase LC is also another commonly used method where, in addition to charge–charge interactions, hydrophobic interactions mainly govern the retention and separation mechanism, making this technique useful for detecting small chemical changes on the full-length sequence. Mixed-mode chromatography, consisting of both reversed-phase and ion-exchange separation modes, provides additional benefits where a single column and method can be used for complete oligonucleotide analysis. Finally, prelimi-nary studies suggest that oligonucleotide-based chromatographic separation may be

another useful method for selective sepa-ration and purification of these important compounds.

References

(1) L.M. Jarvis, Chem. & Eng. News 87(36), 18–27

(2009).

(2) M.A. Zenkova, D.V. Pyshnyi, W.A. Zalloum,

M.B. Gebrezgiabher, and E.V. Bichenkova, Int.

Drug Discovery 6, 28–32 (2011).

(3) D. Bumcrot, M. Manoharan, V. Koteliansky,

and D.W.Y. Sah, Nature Chem. Biol. 2(12),

711–719 (2006).

(4) A. Persidis, Nat. Biotechnol. 17, 403–404

(1999).

(5) S.T. Crooke, Antisense Nuc. Acid Drug Dev.

8(2), 115–122 (1998).

(6) T. Tuschl, Chembiochem. 2, 239–245 (2001).

(7) D. Cejka, D. Losert, and V. Wacheck, Clin. Sci.

110, 47–58 (2006).

(8) E. Song, S.K. Lee, J. Wang, N. Ince, N. Ouy-

ang, J. Min, J. Chen, P. Shankar, and J. Lieber-

man, Nature Med. 9(3), 347–351 (2003).

(9) K.V. Morris, S.W. Chan, D.E. Jacobsen, and

D.J. Looney, Science 305(5688), 1289–1292

(2004).

(10) B.L. Davidson and P.B. McCray, Jr., Nature

Rev. Genetics 12, 329–340 (2011).

(11) M. Manoharan, Curr. Opin. Chem. Biol. 8(6),

570–579 (2004).

(12) J.K. Watts, G.F. Deleavey, and M.J. Damha,

Drug Discovery Today 13(19/20), 842–855

(2008).

(13) F. Czauderna, M. Fechtner, S. Dames, H.

Aygun, A. Klippel, G.J. Pronk, K. Giese, and J.

Kaufmann, Nuc. Acids Res. 31(11), 2705–2716

(2003).

(14) S.L. Beaucage and M.H. Caruthers, Tetrahe-

dron Lett. 22(20), 1859–1862 (1981).

(15) D. Capaldi, K. Ackley, D. Brooks, J. Carmody,

K. Draper, R. Kambhampati, M. Kretschmer,

D. Levin, J. McArdle, B. Noll, R. Raghavachari,

I. Roymoulik, B.P. Sharma, R. Thurmer, and

F. Wincott, Drug Inf. J. 46, 611–626 (2012).

(16) S.P. Waghmare, P. Pousinis, D.P. Hornby, and

M.J. Dickman, J. Chromatogr. A 1216, 1377–

1382 (2009).

(17) M.B. Beverly, Mass Spectrom. Rev. 30, 979–

998 (2011).

(18) A. Andrus, Methods: A companion to Methods in

Enzymol. 4, 213–226 (1992).

(19) F.H.M. van Dinther, R.W. Bally, and T.P.G.

van Sommeren, J. Chromatogr. A 817, 273–279

(1998).

(20) K. Cook and J. Thayer, Bioanalysis 3(10),

1109–1120 (2011).

(21) M. Dizdaroglu and W. Hermes, J. Chromatogr.

171, 321–330 (1979).

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90

80

70

60

50

40

30

20

10

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6.0

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4.0

3.0

2.0

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5.5

4.5

3.5

2.5

1.5

0.5

0.0

1.0

(a)

(b)

Resp

on

se (

mA

U)

Resp

on

se (

mA

U)









N-15: 5’-ACCAAU-3’N-14: 5’-UACCAAU-3’N-13: 5’-AUACCAAU-3’N-12: 5’-AAUACCAAU-3’N-11: 5’-GAAUACCAAU-3’N-10: 5’-UGAAUACCAAU-3’N-9: 5’-CUGAAUACCAAU-3’N-8: 5’-ACUGAAUACCAAU-3’N-7: 5’-CACUGAAUACCAAU-3’N-6: 5’-ACACUGAAUACCAAU-3’N-5: 5’-CACACUGAAUACCAAU-3’N-4: 5’-UCACACUGAAUACCAAU-3’N-3: 5’-AUCACACUGAAUACCAAU-3’N-2: 5’-CAUCACACUGAAUACCAAU-3’N-1: 5’-UCAUCACACUGAAUACCAAU-3’21-mer: 5’-GUCAUCACACUGAAUACCAAU-3’

Figure 7: Chromatographic separation of RNA oligonucleotide samples by mixed mode chromatography using a Scherzo SM-C18 column with a sodium chloride gradi-ent: (a) Separation of N–x deletion series, (b) separation of “base-�ip” isomer stan-dards. Mobile-phase A: 100 mM Tris (pH 7.4) in water; mobile-phase B: 90:10 (v/v) 2 M sodium chloride in 100 mM Tris (pH 7.4) water–acetonitrile; linear gradient: 50–85% B over 35 min; �ow rate: 1 mL/min; column temperature: 50 °C; injection volume: 3 µL; detection: UV absorbance at 260 nm. Adapted with permission from reference 59.


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Mirlinda Bibais an Associate Principal Scientist in the Analytical Chemistry Department at Merck in Rahway, New Jersey. She is also a part-time PhD student at Drexel University in Philadelphia, Pennsylvania, studying under the direction of Professor Joe Foley and Dr. Chris Welch. Her research focuses on analysis and separation of short RNA oligonucleotides by different liquid chromatography techniques.

(22) J.H. van Boom and J.F.M. de Rooy, J. Chro-

matogr. 131, 169-177 (1977).

(23) W. Haupt and A. Pingoud, J. Chromatogr. 260,

419–427 (1983).

(24) W. Jost and K.K. Unger, J. Chromatogr. 185,

403–412 (1979).

(25) J.D. Pearson and F.E. Regnier, J. Chromatogr.

255, 137–149 (1983).

(26) T.G. Lawson, F.E. Regnier, and H.L. Weith,

Anal. Biochem. 133, 85–93 (1983).

(27) R.R. Drager and F.E. Regnier, Anal. Biochem.

145, 47–56 (1985).

(28) H.A. Claessens, M.A. van Straten, and J.J.

Kirkland, J. Chromatogr. A 728, 259–270

(1996).

(29) C. McNeff and P.W. Carr, Anal. Chem. 67,

2350–2353 (1995).

(30) Y. Kato, T. Kitamura, A. Mitsui, and T. Hashi-

moto, J. Chromatogr. 398, 327–334 (1987).

(31) Y. Kato, T. Kitamura, A. Mitsui, Y. Yamasaki,

and T. Hashimoto, J. Chromatogr. 447, 212–

220 (1988).

(32) J.R. Thayer, V. Barreto, S. Rao, and C. Pohl,

Anal. Biochem. 338, 39–47 (2005).

(33) A. Sabarudin, J. Huang, S. Shu, S. Sakagawa,

and T. Umemura, Anal. Chim. Acta 736, 108–

114 (2012).

(34) M. Buncek, V. Backovska, S. Holasova, H.

Radilova, M. Safarova, F. Kunc, and R. Haluza,

Chromatographia 62, 263–269 (2005).

(35) D.G. Volkmann, Anal. Chem. Progress 126,

51–69 (1984).

(36) C. Horvath, W. Melander, I. Molnar, and

P. Molnar, Anal. Chem. 49, 2295–2305

(1977).

(37) R.P.W. Scott and P. Kucera, J. Chromatogr. 175,

51–63 (1979).

(38) P.J. Oefner, J. Chromatogr. B 739, 345–355

(2000).

(39) C.G. Huber, P.J. Oefner, and G.K. Bonn, Anal.

Biochem. 212, 351–358 (1993).

(40) S.M. McCarthy, M. Gilar, and J. Gebler, Anal.

Biochem. 390, 181–188 (2009).

(41) H. Oberacher, A. Krajete, W. Parson, and C.G.

Huber, J. Chromatogr. A 893, 23–35 (2000).

(42) M. Gilar, K.J. Fountain, Y. Budman, U.D.

Neue, K.R. Yardley, P.D. Rainville, R.J. Rus-

sell II, and J.C. Gebler, J. Chromatogr. A 958,

167–182 (2002).

(43) A. Apffel, J.A. Chakel, S. Fischer, K. Lichten-

walter, and W.S. Hancock, Anal. Chem. 69,

1320–1325 (1997).

(44) M. Gilar, Anal. Biochem. 298, 196–206 (2001).

(45) S. Li, D.D. Lu, Y.L. Zhang, and S.Q. Wang,

Chromatographia 72, 215–223 (2010).

(46) B. Noll, S. Seiffert, H.P. Vornlocher, and

I. Roehl, J. Chromatogr. A 1218, 5609–5617

(2011).

(47) K.J. Fountain, M. Gilar, and J.C. Gebler, Rapid

Commun. Mass Spectrom. 17, 646–653 (2003).

(48) V.B. Ivleva, Y.Q. Yu, and M. Gilar, Rapid Com-

mun. Mass Spectrom. 24, 2631–2640 (2010).

(49) M. Beverly, K. Hartsough, L. Machemer, P.

Pavco, and J. Lockridge, J. Chromatogr. B 835,

62–70 (2006).

(50) P. Deng, X. Chen, G. Zhang, and D. Zhong,

J. Pharm. Biomed. Anal. 52, 571–579 (2010).

(51) C.G. Huber, P. J. Oefner, E. Preuss, and G. K.

Bonn, Nuc. Acids Res. 21(5), 1061–1066 (1993).

(52) C.G. Huber, P.J. Oefner, and G.K. Bonn, Anal.

Chem. 67, 578–585 (1995).

(53) C.G. Huber, E. Stimpf, P.J. Oefner, and G.K.

Bonn, LCGC North Am. 14(2), 114–127 (1996).

(54) M. Biba, C.J. Welch, J.P. Foley, B. Mao, E.

Vazquez, and R.A. Arvary, J. Pharm. Biomed.

Anal. 72, 25–32 (2013).

(55) T. Kremmer, M. Boldizsar, and L. Holczinger,

J. Chromatogr. 415, 53–63 (1987).

(56) L.W. McLaughlin, J. Chromatogr. 418, 51–72

(1987).

(57) R. Bischoff and L.W. McLaughlin, Anal. Bio-

chem. 151, 526–533 (1985).

(58) L.W. McLaughlin, Chem. Rev. 89, 309–319

(1989).

(59) M. Biba, E. Jiang, B. Mao, D. Zewge, J.P. Foley,

and C.J. Welch, J. Chromatogr. A 1304, 69–77

(2013).

(60) S. Okamura, F. Crane, H.A. Messner, and

T.W. Mak, J. Biol. Chem. 253(11), 3765–3767

(1978).

(61) R.H. Hovhannisyan and R.P. Carstens, Bio-

Techniques 46, 95–98 (2009).

(62) E.A. Smith, M. Kyo, H. Kumasawa, K. Naka-

tani, I. Saito, and R.M. Corn, J. Am. Chem. Soc.

124, 6810–6811 (2002).

(63) F. Seela and S. Budow, Mol. BioSyst. 4, 232–

245 (2008).

(64) M. Biba, C.J. Welch, and J.P. Foley, unpub-

lished data (2013).

(65) Y. Aiba, Y. Hamano, W. Kameshima, Y. Araki,

T. Wada, A. Accetta, S. Sforza, R. Corradini, R.

Marchelli, and M. Komiyama, Org. & Biomol.

Chem. 11(32), 5233–5238 (2013).



Bing Maois currently Director, Analytical Chemistry within Process and Analytical Chemistry Department at Merck Research Laboratories in Rahway. He has more than 17 years of experience at Merck with analytical characterization and development to support pharmaceutical small molecule, peptide, and oligonucleotide drug substance manufacturing process development and optimization.

Christopher J. Welchis a science lead for analytical chemistry within the Process and Analytical Chemistry area at Merck Research Laboratories in Rahway. Chris also co-chairs the New Technologies Review and Licensing Committee (NT-RLC), the organization that oversees identification, acquisition, and evaluation of new technologies of potential value to Merck Research Laboratories. Chris also co-chairs the MRL Postdoctoral Research Fellows Program.

Joe P. Foleyis Professor of Chemistry and Associate Department Head at Drexel University, and a lifetime member of the Chromatography Forum of the Delaware Valley (CFDV). He received his PhD in Chemistry from the University of Florida, and followed it with a two-year NRC postdoc at NIST. His research interests are in the fundamental and applied aspects of analytical chemistry and separation science, and he has authored or co-authored more than 110 articles, book chapters, reviews, and one patent pertaining to pressure- and voltage-driven liquid-phase chiral and achiral separations (that is, HPLC, UHPLC, SFC, and CE/EKC).

Direct correspondence to: [email protected] �


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