Comparison of Silica-Based Monoliths and Organic Monoliths

Một phần của tài liệu Handbook of HPLC Second Edition (Trang 56 - 713)

1.6 Chromatographic Applications of Organic and Inorganic Monoliths

1.6.3 Comparison of Silica-Based Monoliths and Organic Monoliths

The strict distinction between silica-based monolithic columns and polymeric columns in most papers and reviews is likely attributed to obvious differences with respect to preparation chemistry and consequently fabrication procedure. Because of the simplicity of the fabrication of organic monoliths, a huge number of monomer chemistry and a wide variety of different columns for a multitude of promising applications have been described in literature. Even the basic synthesis pro- tocol of organic polymers seems to be straightforward, adjusting (fine tuning) the porous properties in order to yield the desired chromatographic characteristics toward the target analytes is criti- cal and demands experience as well as know-how and scientific patience. Silica monoliths, on the other hand, are more difficult and challenging to prepare, in particular with respect to the sensitive

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FIGure 1.15 Separation of the 16 EPA priority pollutants PAHs with ODS column using an acetonitrile:water 70:30 (v/v) solution as mobile phase. Thiourea was used as tM standard. Detection performed at 254 nm and 30°C.

PAHs: 1, naphthalene; 2, acenaphtylene; 3, fluorene; 4, acenaphthene; 5, phenanthrene; 6, anthracene; 7, fluo- ranthene; 8, pyrene; 9, chrysene; 10, benz(a)anthracene; 11, benzo(b)fluoranthene; 12, benzo(k)fluoranthene; 13, benzo(a)pyrene; 14, dibenz(a,h)anthracene; 15, indeno(1,2,3-cd)pyrene; and 16, benzo(g,h,i)perylene). (Reprinted from Nunez, O. et al., J. Chromatogr. A, 1175, 7, 2007. Copyright 2007, with permission from Elsevier.)

“ageing” procedures, which have to be employed after the actual polymerization in order to obtain the desired mesoporous structure and pore-size distribution. Few groups reported on successful attempts for reproducing the preparation methods, initially introduced by Nakanishi et al. Up to now, literature on silica monoliths is mainly restricted to bare silica or C18 derivatized silica rods, lacking the huge range of different chemistries offered by their organic counterpart.

The second distinction between organic and inorganic (silica) monolith refer to the typical and preferred column dimension. Even if mechanically stable organic monolithic materials have been fabricated in almost every column dimension, ranging from nanocolumns (20 μm I.D.), over capil- lary columns to conventional column (2–8 mm I.D.) and even to the preparative format, their main focus with respect chromatographic application is put on capillary column with an I.D. > 200 μm.

This might be explained by the fact that free radical polymerization, which is a strongly exothermic reaction, creates a radial temperature gradient across the column, which is the more pronounced the larger the diameter of the column mold is [58]. This temperature gradient, in turn, influences the rate of polymerization and thus causes inhomogeneity of the resulting monolith porosity, which are closely associated with decreased column performance. In fact, a direct comparison of conventional HPLC columns (3 mm I.D.), based on a monolithic styrene network, has revealed a considerable loss in efficient than their capillary counterparts (200 μm I.D.) [140]. Silica monoliths, on the other hand, are mainly produced as conventional sized HPLC columns, even if recently Merck commercialized a silica monolith in a 100 μm fused silica capillary. However, narrow pore and capillary columns within a diameter range of 100–500 μm are scarcely reported in literature. This is attributed to the extensive shrinkage of the monolithic skeleton during the drying process, which demands for a cladding (leakproof fit into a column housing) procedure after polymerization. In case the silica rod is prepared in a sufficiently narrow (I.D. < 100 μm) silica tube, the gel remains glued to the wall. As the cladding process is from a technical point of view exceedingly difficult to realize for capillary columns up to 500 μm, miniaturized column between 100 and 500 μm I.D. cannot be prepared following the classical sol–gel approach. However, recently, alternative polymerization procedures have been introduced that allow the preparation of narrow pore and capillary silica monoliths without wall detachment [213].

Finally, silica and polymer monoliths significantly differ in their preferred field of application.

While organic monoliths numerous times proved to enable highly efficient and swift resolution of large biomolecules, silica monoliths focus on the application of monolithic stationary phases by providing powerful and promising result for fast and high-resolution separation of low-molecular- weight compounds. The characteristic bimodal pore size distribution of silica monoliths establishes separation performance for small molecules, ascribed due to a high fraction of accessible mesopo- res, whereas the monomodal macropore distribution of typical organic monoliths comply with the requirements of macromolecule chromatography, mainly by reducing resistance to mass transfer by substituting analyte diffusion by solvent convention.

However, silica monoliths and organic polymers both exhibit very advantageous chromatographic characteristics: enhanced mass transfer characteristics, high reproducibility, and versatile surface chemistry, which make monolithic column attractive for a variety of forward-looking applications.

reFerenCes

1. H. Staudinger, E. Huseman. Berichte der Deutschen Chemischen Gesellschaft 68, 1618–1634, 1935.

2. G.F. D’Alelio. U.S. Patent 2,366,007, 1945.

3. K.W. Pepper. J. Appl. Chem. 1, 124–132, 1951.

4. I.M. Abrams. J. Ind. Eng. Chem. 48, 1469–1472, 1956.

5. O. Okay. Prog. Polym. Sci. 25, 711–779, 2000.

6. K. Dusek. Chem. Prumysl. 11, 439–443, 1961.

7. R. Kunin, E. Meitzner, N. Bortnick. J. Am. Chem. Soc. 84, 305–306, 1962.

8. J.R. Millar, D.G. Smith, W.E. Marr, T.R.E. Kressman. J. Chem. Soc. 2779–2784, 1963.

9. J. Malinsky, J. Rahm, F. Krska, J. Seidl. Chem. Prumysl. 13, 386, 1963.

10. F. Svec, F.M.F. Fréchet. Anal. Chem. 64, 820–822, 1992.

11. D.L. Mould, R.L.M. Synge. Analyst 77, 964–970, 1952.

12. D.L. Mould, R.L.M. Synge. Biochem. J. 58, 571–585, 1954.

13. M. Kubin, P. Spacek, R. Chromekec. Coll. Czechosl. Chem. Commun. 32, 3881–3887, 1967.

14. H. Schnecko, O. Bieber. Chromatographia 4, 109–112, 1971.

15. L.C. Hansen, R.E. Sievers. J. Chromatogr. 99, 123–133, 1974.

16. F.D. Hileman, R.E. Sievers, G.G. Hess, W.D. Ross. Anal. Chem. 45, 1126–1130, 1973.

17. T.R. Lynn, D.R. Rushneck, A.R. Cooper. J. Chromatogr. Sci. 12, 76–79, 1974.

18. S. Hjérten, J.L. Liao, R. Zhang. J. Chromatogr. 473, 273–275, 1989.

19. J.L. Liao, R. Zang, S. Hjérten. J. Chromatogr. 586, 21–26, 1991.

20. B.G. Belenkii, A.M. Podkladenko, O.I. Kurenbin, V.G. Maltsev, D.G. Nasledov, S.A. Trushin.

J. Chromatogr. A 645, 1–15, 1993.

21. T.B. Tennikova, B.G. Belenkii, F. Svec. J. Liquid Chromatogr. 13, 63–70, 1990.

22. F. Svec, T.B. Tennikova. J. Bioact. Compat. Polym. 6, 393–405, 1991.

23. T.B. Tennikova, F. Svec. J. Chromatogr. 646, 279–288, 1993.

24. Q.C. Wang, F. Svec, J.M.J. Fréchet. Anal. Chem. 65, 2243–2248, 1993.

25. E.C. Peters, F. Svec, J.M.J. Fréchet. Adv. Mater. 11, 1169–1181, 1999.

26. F. Svec. J. Sep. Sci. 27, 1419–1430, 2004.

27. F. Svec, E.C. Peters, D. Sykora, J.M.J. Fréchet. J. Chromatogr. A 887, 3–29, 2000.

28. E. Klodzin′ska, D. Moravcova, P. Jandera, B. Buszewski. J. Chromatogr. A 1109, 51–59, 2006.

29. F. Svec, C.G. Huber. Anal. Chem. 78, A2101–A2107, 2006.

30. B.G. Belenkii. Russ. J. Bioorg. Chem. 32, 323–332, 2006.

31. F. Svec, J.M.J. Fréchet. Ind. Eng. Chem. Res. 38, 34–48, 1999.

32. K. Štulik, V. Pacakova, J. Suchankova, P. Coufal. J. Chromatogr. B 841, 79–87, 2006.

33. H. Minakuchi, N. Nakanishi, N. Soga, N. Ishizuka, N. Tanaka. Anal. Chem. 68, 3498–3501, 1996.

34. S.M. Fields. Anal. Chem. 68, 2709–2712, 1996.

35. J.F. Keenedy, G.O. Phillips, P.A. Williams. Cellulosics: Materials for Selective Separations and Other Technologies. Horwood, New York, 1993.

36. F. Sinner, M.R. Buchmeiser. Macromolecules 33, 5777–5786, 2000.

37. B. Mayr, R. Tessadri, E. Post, M.R. Buchmeiser. Anal. Chem. 73, 4071–4078, 2001.

38. B. Mayr, G. Hửlzl, K. Eder, M.R. Buchmeiser, C.G. Huber. Anal. Chem. 74, 6080–6087, 2002.

39. S. Lubbad, M.R. Buchmeiser. Macromol. Rapid Commun. 23, 617–612, 2002.

40. K. Hosoya, N. Hira, K. Yamamoto, M. Nishimura, N. Tanaka. Anal. Chem. 78, 5729–5735, 2006.

41. Q.C. Wang, F. Svec, J.M.J. Fréchet. Anal. Chem. 65, 2243, 1993.

42. M. Petro, F. Svec, J.M.J. Fréchet. J. Chromatogr. A 752, 59, 1996.

43. Q.C. Wang, F. Svec, J.M.J. Fréchet. J. Chromatogr. A 669, 230, 1994.

44. A. Premstaller, H. Oberacher, W. Walcher, A.M. Timperio, L. Zolla, J.-P. Chervet, N. Cavusoglu, A. van Dorsselaer, C.G. Huber. Anal. Chem. 73, 2390, 2001.

45. W. Walcher, H. Oberacher, S. Troiani, G. Hửlzl, P. Oefner, L. Zolla, C.G. Huber. J. Chromatogr. B 782, 111, 2002.

46. W. Walcher, H. Toll, A. Ingendoh, C.G. Huber. J. Chromatogr. A 1053, 107, 2004.

47. H. Oberacher, A. Krajete, W. Parson, C.G. Huber. J. Chromatogr. A 893, 23, 2000.

48. H. Oberacher, C.G. Huber. Trend. Anal. Chem. 21, 166, 2002.

49. A. Premstaller, H. Oberacher, C.G. Huber. Anal. Chem. 72, 4386–4393, 2000.

50. G. Hửlzl, H. Oberacher, S. Pitsch, A. Stutz, C.G. Huber. Anal. Chem. 77, 673, 2005.

51. C.G. Huber, A. Krajete. Anal. Chem. 71, 3730, 1999.

52. X. Huang, S. Zhang, G.A. Schultz, J. Henion. Anal. Chem. 74, 2336, 2002.

53. F. Svec. J. Sep. Sci. 27, 747–766, 2004.

54. A. Podgornik, M. Barut, J. Jancar, A. Strancar. J. Chromatogr. A 848, 51, 1999.

55. D. Sykora, F. Svec, J.M.J. Fréchet. J. Chromatogr. A 852, 297, 1999.

56. F. Svec, J.M.J. Fréchet. J. Chromatogr. A 1995, 89, 1995.

57. E. Suarez, B. Paredes, F. Rubiera, M. Rendueles, M.A. Villa-Garcia, J.M. Diaz. Sep. Purif. Technol.

27, 1, 2002.

58. A. Podgornik, M. Barut, A. Strancar, D. Josic, T. Koloini. Anal. Chem. 72, 5693, 2000.

59. M. Barut, A. Podgornik, P. Brne, A. Strancar. J. Sep. Sci. 28, 1876, 2005.

60. J. Urthaler, R. Schlegl, A. Podgornik, A. Strancar, A. Jungbauer, R. Necina. J. Chromatogr. A 1065, 93, 2005.

61. M. Bencina, A. Podgornik, A. Strancar. J. Sep. Sci. 27, 801, 2004.

62. C. Viklund, F. Svec, J.M.J. Fréchet, K. Irgum. Biotechnol. Prog. 13, 597, 1997.

63. Q. Luo, H. Zou, X. Xiao, Z. Guo, L. Kong, X. Mao. J. Chromatogr. A 926, 255, 2001.

64. M. Petro, F. Svec, J.M.J. Fréchet. Biotechnol. Bioeng. 49, 355, 1996.

65. S. Xie, F. Svec, J.M.J. Fréchet. Biotechnol. Bioeng. 62, 30, 1999.

66. S. Hjertén, J.-L. Liao, R. Zhang. J. Chromatogr. 473, 273, 1989.

67. J.-L. Liao, S. Zhang, S. Hjertén. J. Chromatogr. 586, 21, 1991.

68. Y.-M. Li, J.-L. Liao, K. Nakazato, J. Mohammad, L. Terenius, S. Hjertén. Anal. Biochem. 223, 153, 1994.

69. F.M. Sinner, M.R. Buchmeiser. Angew. Chem. Int. Ed. 39, 1433, 2000.

70. C. Gatschelhofer, C. Magnes, T.R. Pieber, M.R. Buchmeiser, F.M. Sinner, J. Chromatogr. A 1090, 81, 2005.

71. S. Lubbad, B. Mayr, C.G. Huber, M.R. Buchmeiser, J. Chromatogr. A 959, 121, 2002.

72. B. Mayr, R. Tessadri, E. Post, M.R. Buchmeiser. Anal. Chem. 73, 4071, 2001.

73. L.A. Errede. Polym. Preprints 26, 77, 1985.

74. L.A. Errede. J. Appl. Polym. Sci. 31, 1746–1761, 1986.

75. J. Vidicˇ, A. Podgornik, A. Štrancar. J. Chromatogr. A 1065, 51–58, 2005.

76. K. Nakanishi, N. Soga. J. Am. Ceram. Soc. 74, 2518, 1991.

77. K. Nakanishi, N. Soga. J. Non-Cryst. Solids 139, 1, 1992.

78. K. Nakanishi, N. Soga. J. Non-Cryst. Solids 139, 14, 1992.

79. K. Nakanishi, N. Soga, Inorganic porous column, Japan Patent 5-200,392, 1993.

80. K. Nakanishi, N. Soga, Production of inorganic porous body, Japan Patent 5-208,642, 1993.

81. K. Nakanishi, N. Soga, Inorganic porous material and process for making same, U.S. Patent 5,624,875, 1997.

82. K. Cabrera, G. Sọttler, G. Wieland, Trennmittel (separator), European patent EP 0,686,258 b1, 1994.

83. N. Tanaka, H. Kobayashi, N. Ishizuka, H. Minakushi, K. Nakanishi, K. Hosoya, T. Itegami. J. Chromatogr.

A 965, 35, 2002.

84. T. Ikegami, H. Kujita, K. Horie, K. Hosoya, N. Tanaka. Anal. Bional. Chem. 386, 578, 2006.

85. S. Wienkoop, M. Glinski, N. Tanaka, V. Tolstikof, O. Fiehn, W. Weckwerth. Rapid Commun. Mass Spectrom. 18, 643, 2004.

86. M. Kato, K. Inuzuka, K. Sakai-Kato, T. Toyo’oka. Anal. Chem. 77, 1813, 2005.

87. A.-M. Siouffi. J. Chromatogr. A 1000, 801, 2003.

88. A. Vegvari. J. Chromatogr. A 1079, 50, 2005.

89. N. Ishizuka, H. Minakuchi, K. Nakanishi, K. Hirao, N. Tanaka, Colloids Surf. A 187–188, 273, 2001.

90. H. Saito, K. Nakanishi, K. Hirao, H. Jinnai. J. Chromatogr. A 1119, 95, 2006.

91. K. Nakanishi, H. Minakuchi, N. Soga, N. Tanaka. J. Sol–Gel Sci. Technol. 13, 163, 1998.

92. C. Yang, T. Ikegami, T. Hara, N. Tanaka. J. Chromatogr. A 1130, 175, 2006.

93. O. Nunez, T. Ikegami, K. Miyamoto, N. Tanaka. J. Chromatogr. A 1175, 7–15, 2007.

94. L.A. Colon, D.C. Hoth, Group IV Metal Oxide Monolithic Columns, PCT Int. Appl. WO200509, 1972, 2005.

95. D.C. Hoth, J.G. Rivera, L.A. Colón. J. Chromatogr. A 1079, 392, 2005.

96. J. Randon, S. Huguet, A. Piram, G. Puy, C. Demesmay, J.-L. Rocca. J. Chromatogr. A 1109, 19, 2006.

97. A. Taguchi, J.-H. Smatt, M. Linden. Adv. Mater. 15, 1209, 2003.

98. C. Liang, S. Dai, G. Guiochon. Anal. Chem. 75, 4904, 2003.

99. C. Liang, Synthesis and applications of monolithic HPLC columns, PhD Thesis, University of Tennessee, Knoxville, TN, 2005.

100. E.C. Peters, C. Ericson. In: Monolithic Materials, J. Chromatogr. Libr., Vol. 67. Elsevier, Amsterdam, the Netherlands, 2003.

101. J. Seidl, J. Malinsky, K. Dusek, W. Heitz. Fortschritte der Hochpolymeren-Forschung, 5, 113–213, 1967.

102. A. Guyot, M. Bartholin. Prog. Polym. Sci. 8, 277–331, 1982.

103. K.A. Kun, R. Kunin. J. Polym. Sci. 6, 2689–2701, 1968.

104. W.L. Sederel, G.J. DeJong. J. Appl. Polym. Sci. 17, 2835–2846, 1973.

105. C. Viklund, F. Svec, J.M.J. Fréchet, K. Irgum. Chem. Mater. 8, 744–750, 1996.

106. S. Xie, F. Svec, J.M.J. Fréchet. J. Polym. Sci. A 35, 1013–1021, 1997.

107. C. Viklund, E. Ponten, B. Glad, K. Irgum, P. Hửrstedt, F. Svec. Chem. Mater. 9, 463–471, 1997.

108. F. Svec, J.M.J. Fréchet. Chem. Mater. 7, 707–715, 1995.

109. F. Svec, J.M.J. Fréchet. Macromolecules 28, 7580–7582, 1995.

110. J. Brandrup, E.H. Immergut. Polymer Handbook. Wiley, New York, 1989.

111. G. Bonn, S. Lubbad, L. Trojer. U.S. Patent 144,971 A1, 2007.

112. L. Trojer, G. Stecher, I. Feuerstein, G.K. Bonn. Rapid Commun. Mass Spectrom. 19, 3398–3404, 2005.

113. L. Trojer, G. Stecher, I. Feuerstein, S. Lubbad, G.K. Bonn. J. Chromatogr. A 1079, 197–207, 2005.

114. K.S.W. Sing, D.H. Everett, R.A.W. Haul, L. Moscou, R.A. Pierotti, J. Rouquerol, T. Siemieniewska. Pure Appl. Chem. 57, 603–619, 1985.

115. E.W. Washburn. Proc. Natl. Acad. Sci. U.S.A. 7, 115, 1921.

116. E.W. Washburn. Phys. Rev. 17, 273, 1921.

117. H.M. Rootare, C.F. Prenzlow. J. Phys. Chem. 71, 2733, 1967.

118. P.H. Emmett, T.W. DeWitt. J. Am. Chem. Soc. 65, 1253, 1943.

119. I. Halász, K. Martin. Angew. Chem. 90, 954, 1978.

120. M.E. Van Kreveld, N. Van den Hoed. J. Chromatogr. 83, 111–124, 1973.

121. H.W. Siesler., Y. Ozaki, S. Kawata, H.M. Heise, Near-Infrared Spectroscopy—Principles, Instruments, Applications. Wiley-VCH, Weinheim, Germany, 2002.

122. J. Workman, L. Weyer, Practical Guide to Interpretive Near-infrared Spectroscopy. Taylor & Francis, Boca Raton, FL, 2007.

123. D.A. Burns, E.W. Ciurczak, Handbook of Near-Infrared Analysis. CRC Press, Boca Raton, FL, 2001.

124. B.J. Berne, R. Pecora, Dynamic Light Scattering. Dover Publications, Inc. New York, 2000.

125. H.C. van de Hulst, Light Scattering by Small Particles. Dover Publications, Inc., New York, 1981.

126. D.J. Dahm, K.D. Dahm, K.H. Norris. J. Near Infrared Spectrosc. 10(1), 1–13, 2002.

127. C.W. Huck, R. Ohmacht, Z. Szabo, G.K. Bonn. J. Near Infrared Spectrosc. 14, 51–57, 2006.

128. D.J. Dahm, K.D. Dahm, Interpreting Diffuse Reflectance and Transmittance. IM Publications, Chichester, U.K., 2007.

129. N. Heigl, C.H. Petter, M. Rainer, M. Najam-ul-Haq, R.M. Vallant, R. Bakry, G.K. Bonn, C.W. Huck.

J. Near Infrared Spectrosc. 15(5), 269–282, 2007.

130. C.W. Huck, N. Heigl, M. Najam-ul-Haq, M. Rainer, R.M. Vallant, G.K. Bonn. Open Anal. Chem. J.

1, 21–27, 2007.

131. C.A. Cramers, J.A. Rijks, C.P.M. Schutjes. Chromatographia 14, 439, 1981.

132. P.A. Bristow, J.H. Knox. Chromatographia 10, 279, 1977.

133. F. Nevejans, M. Verzele. J. Chromatogr. 350, 145, 1985.

134. M. Petro, F. Svec, I. Gitsov, J.M.J. Fréchet. Anal. Chem. 68, 315–321, 1996.

135. Q.C. Wang, F. Svec, J.M.J. Fréchet. J. Chromatogr. A 669, 230–235, 1994.

136. I. Gusev, X. Huang, Cs. Horvath. J. Chromatogr. A 855, 273–290, 1999.

137. W. Walcher, H. Oberacher, S. Troiani, G. Hửlzl, P. Oefner, L. Zolla, C.G. Huber. J. Chromatogr. B 782, 111–125, 2002.

138. Q.C. Wang, F. Svec, J.M.J. Fréchet. Anal. Chem. 67, 670–674, 1995.

139. L. Trojer, S.H. Lubbad, C.P. Bisjak, G.K. Bonn. J. Chromatogr. A 1117, 56–66, 2006.

140. L. Trojer, S.H. Lubbad, C.P. Bisjak, W. Wieder, G.K. Bonn. J. Chromatogr. A 1146, 216–224, 2007.

141. F. Svec, J.M.J. Fréchet. J. Chromatogr. A 702, 89–95, 1995.

142. C. Viklund, F. Svec, J.M.J. Fréchet. Biotechnol. Prog. 13, 597–600, 1997.

143. D. Sykora, F. Svec, J.M.J. Fréchet. J. Chromatogr. A 852, 297–304, 1999.

144. Q. Luo, H. Zou, X. Xiao, Z. Guo, L. Kong, X. Mao. J. Chromatogr. A 926, 255–264, 2001.

145. Y. Ueki, T. Umemura, J. Li, T. Odake, K. Tsunoda. Anal. Chem. 76, 7007–7012, 2004.

146. E.C. Peters, M. Petro, F. Svec, J.M.J. Fréchet. Anal. Chem. 69, 3646–3649, 1997.

147. E.C. Peters, M. Petro, F. Svec, J.M.J. Fréchet. Anal. Chem. 70, 2296–2302, 1998.

148. G. Ping, L. Zhang, L. Zhang, W. Zhang, P. Schmitt-Kopplin, A. Kettrup, Y. Zhang. J. Chromatogr. A 1035, 265–270, 2004.

149. D. Lee, F. Svec, J.M.J. Fréchet. J. Chromatogr. A 1051, 53–60, 2004.

150. L. Geiser, S. Eeltnik, F. Svec, J.M.J. Fréchet. J. Chromatogr. A 1140, 140–146, 2007.

151. C. Yu, M. Xu, F. Svec, J.M.J. Fréchet. J. Polym. Sci. A 40, 755–769, 2002.

152. P. Coufal, M. Cˇihak, J. Suchankova, E. Tesarˇova, Z. Bosanova, K. Štulk. J. Chromatogr. A 946, 99–106, 2002.

153. D. Moravcova, P. Jandera, J. Urban, J. Planeta. J. Sep. Sci. 26, 1005–1016, 2003.

154. P. Holdšvendova, P. Coufal, J. Suchankova, E. Tesarˇova, Z. Bosakova. J. Sep. Sci. 26, 1623–1628, 2003.

155. T. Rohr, C. Yu, M.H. Davey, F. Svec, J.M.J. Fréchet. Electrophoresis 22, 3959–3967, 2001.

156. C. Yu, M.H. Davey, F. Svec, J.M.J. Fréchet. Anal. Chem. 73, 5088–5096, 2001.

157. L. Uzun, R. Say, A. Denizli. React. Funct. Polym. 64, 93–102, 2005.

158. P. Hemstrửm, A. Nordborg, K. Irgum, F. Svec, J.M.J. Frechet. J. Sep. Sci. 29, 25–32, 2006.

159. D.S. Peterson, T. Rohr, F. Svec, J.M.J. Fréchet. Anal. Chem. 74, 4081–4088, 2002.

160. S. Xie, F. Svec, J.M.J. Fréchet. Biotechnol. Bioeng. 62, 30–35, 1999.

161. C. Viklund, A. Sjogren, K. Irgum. Anal. Chem. 73, 444–452, 2000.

162. C. Viklund, K. Irgum. Macromolecules 33, 2539–2544, 2000.

163. Z. Jiang, N.W. Smith, P.D. Ferguson, M.R. Taylor. Anal. Chem. 79, 1243–1250, 2007.

164. B. Gu, J.M. Armenta, M.L. Lee. J. Chromatogr. A 1079, 382–391, 2005.

165. L.J. Sondergeld, M.E. Bush, A. Bellinger, M.M. Bushey. J. Chromatogr. A 1004, 155–165, 2003.

166. B.L. Waguespack, S.A. Slade, A. Hodges, M.E. Bush, L.J. Sondergeld, M.M. Bushey. J. Chromatogr. A 1078, 171–180, 2005.

167. M. Bedair, Z. El Rassi. Electrophoresis 23, 2938–2948, 2002.

168. S.M. Ngola, Y. Fintschenko, W.-Y. Choi, T.J. Shepodd. Anal. Chem. 73, 849–856, 2001.

169. R. Shediac, S.M. Ngola, D.J. Throckmorton, D.S. Anex, T.J. Shepodd, A.K. Singh. J. Chromatogr. A 925, 251–263, 2001.

170. O. Kornyšova, A. Maruška, P.K. Owens, M. Erickson. J. Chromatogr. A 1071, 171–178, 2005.

171. C.P. Bisjak, S.H. Lubbad, L. Trojer, G.K. Bonn. J. Chromatogr. A 1147, 46–52, 2007.

172. C.P. Bisjak, L. Trojer, S.H. Lubbad, W. Wieder, G.K. Bonn. J. Chromatogr. A 1154, 269–276, 2007.

173. S. Xie, F. Svec, J.M.J. Fréchet. J. Chromatogr. A 775, 65–72, 1997.

174. D. Hoegger, R. Freitag. J. Chromatogr. A 914, 211–222, 2001.

175. D. Hoegger, R. Freitag. Electrophoresis 24, 2958–2972, 2003.

176. F.M. Plieva, J. Andersson, I.Y. Galaev, B. Mattiasson. J. Sep. Sci. 27, 828–836, 2004.

177. P. Arvisson, F.M. Plieva, V.I. Lozinsky, I.Y. Galaev, B. Mattiasson. J. Chromatogr. A 986, 275–290, 2003.

178. F.M. Plieva, I.N. Savina, S. Deraz, J. Andersson, I.Y. Galaev, B. Mattiasson. J. Chromatogr. B 807, 129–137, 2004.

179. A. Maruška, C. Ericson, A. Vegvari, S. Hjerten. J. Chromatogr. A 837, 25–33, 1999.

180. S. Xie, R.W. Allington, F. Svec, J.M.J. Fréchet. J. Chromatogr. A 865, 169, 1999.

181. G. Yue, Q. Luo, J. Zhang, S. Wu, B.L. Karger. Anal. Chem. 79, 938–946, 2007.

182. R.M. Vallant, Z. Szabo, L. Trojer, M. Najam-ul-Haq, M. Rainer, C.W. Huck, R. Bakry, G.K. Bonn.

J. Proteome Res. 6, 44–53, 2006.

183. W. Wieder, S.H. Lubbad, L. Trojer, C. Bisjak, G.K. Bonn. J. Chromatogr. A 1191, 253–262, 2008.

184. N. Marti, PhD thesis, Institute for Chemical and Bioengineering, ETH, Zürich, Switzerland, 2007.

185. C.P. Bisjak, R. Bakry, C.W. Huck, G.K. Bonn. Chromatographia 62, 31–36, 2005.

186. S. Hjerten, H.J. Issaq. A Century of Separation Science, Marcel Dekker, Inc., New York, p. 421, 2001.

187. M. Rezeli, F. Kilar, S. Hjerten. J. Chromatogr. A 1109, 100, 2006.

188. J.-L. Liao, Y. Wang, S. Hjerten. Chromatographia 42, 259, 1996.

189. H. Minakuchi, K. Nakanishi, N. Soga, N. Tanaka. J. Chromatogr. A 828, 83, 1998.

190. B. Barroso, D. Lubda, R. Bishoff. J. Proteome Res. 2, 633, 2003.

191. H. Kimura, T. Tanigawa, H. Morisaka, T. Ikegami, K. Hosoya, N. Ishizuka, H. Minakuchi, K. Nakanishi, M. Ueda, K. Cabrera, N. Tanaka. J. Sep. Sci. 27, 897, 2004.

192. L. Rieux, H. Niederlọnder, E. Velpoorte, R. Bischoff. J. Sep. Sci. 28, 1628, 2005.

193. L. Rieux, D. Lubda, H.A.G. Niederlọnder, E. Verpoorte, R. Bischoff. J. Chromatogr. A 1120, 165, 2006.

194. T. Ikegami, K. Horie, J. Jaafar, K. Hosoya, N. Tanaka. Biochem. Biophys. Methods 70, 31, 2007.

195. M. Kato, K. Satai-Kato, H. Jin, K. Kubota, H. Miyano, T. Toyooka, M.T. Dulay, R.N. Zare. Anal. Chem.

76, 1896, 2004.

196. T. Ikegami, T. Hara, H. Kimura, H. Kobayashi, K. Hosoya, K. Cabrera, N. Tanaka. J. Chromatogr. A 1106, 112, 2006.

197. N. Tanaka, H. Kimura, D. Tokuda, K. Hosoya, T. Ikegami, N. Ishizuka, H. Minakuchi, K. Nakanishi, Y. Shintani, M. Furuno, K. Cabrera. Anal. Chem. 76, 1273, 2004.

198. B.W. Pack, D.S. Risley. J. Chromatogr. A 1073, 269, 2005.

199. B. Chankvetadze, C. Yamamoto, M. Kamigaito, N. Tanaka, K. Nakanishi, Y. Okamoto, J. Chromatogr. A 1110, 46, 2006.

200. A.H. Schmidt. J. Chromatogr. A 1073, 377, 2005.

201. L. Jia, N. Tanaka, S. Terabe. J. Chromatogr. A 1053, 71, 2004.

202. S. Brunauer. Physical Adsorption, Vol. 1. Princeton University Press, Princeton, NJ, 1945.

203. S. Brunauer, P.H. Emmet. J. Am. Chem. Soc. 57, 1754–1755, 1935.

204. S. Brunauer, P.H. Emmett, E. Teller. J. Am. Chem. Soc. 60, 309–319, 1938.

205. K.K. Unger, R. Janzen, G. Jilge. Chromatographia 24, 144–154, 1987.

206. H. Oberacher, A. Premstaller, C.G. Huber. J. Chromatogr. A 1030, 201–208, 2004.

207. M. Kele, G. Guiochon. J. Chromatogr. A 960, 19–49, 2002.

208. M. Kele, G. Guiochon. J. Chromatogr. A 830, 41–54, 1999.

209. M. Kele, G. Guiochon. J. Chromatogr. A 830, 55–79, 1999.

210. M. Kele, G. Guiochon. J. Chromatogr. A 855, 423–453, 1999.

211. M. Kele, G. Guiochon. J. Chromatogr. A 869, 181–209, 2000.

212. M. Kele, G. Guiochon. J. Chromatogr. A 913, 89–112, 2001.

213. G. Guiochon. J. Chromatogr. A 1168, 101–168, 2007.

214. M. Motokawa, H. Kobayashi, N. Ishizuka, H. Minakushi, K. Nakanishi, H. Jinnai, K. Hosoya, T. Itegami, N. Tanaka. J. Chromatogr. A 961, 53, 2002.

215. D. Moravcova, P. Jandera, J. Urban, J. Planeta. J. Sep. Sci. 27, 789, 2004.

47

2 Bonded Stationary Phases

Heinz Engelhardt

2.1 IntroduCtIon

High-performance liquid chromatography (HPLC) has become the dominant analytical technique in pharmaceutical, chemical, and food industries, as well as in environmental laboratories, in clini- cal chemistry for therapeutic drug monitoring, and in bioanalysis [1]. The rise of HPLC to the most widely used instrumental analytical systems originates in part from the broad variety of selectivities introduced by the enormous number of stationary phases available and the easy adjustment of selec- tivity by changing the composition of the mobile phase. The classical separation systems based on pure silica or alumina—now called normal phase chromatography—with nonpolar mobile phases would not have provided the variety and the simplicity of separation methods, and the reproduc- ibility, now state-of-the-art in HPLC. The availability of the so-called reversed phases (RPs) based on chemically modified silica, where adsorption is the highest from aqueous solutions, opened for HPLC direct access to aqueous, and hence, bioanalytical systems. These phases are the workhorses in HPLC. Their diversity allows us to select appropriate columns for a wide variety of applications ranging from separations of aromatic hydrocarbons, pharmaceuticals, and pesticides to applications Contents

2.1 Introduction ... 47 2.2 The Base Material: Silica ... 49 2.2.1 Physical Parameters ... 49 2.2.2 Measurement of Physical Parameters ...50 2.2.3 Chemical Properties of Silica ...50 2.2.4 Bonding Technology ... 51 2.2.5 Bonded Phases with Functional Groups ... 53 2.2.6 RP with Shielding Groups (Polar-Embedded RP) ...54 2.2.7 RP with Other Functional Groups ... 55 2.2.8 Alternative Bonding Technologies ... 57 2.2.9 Other Inorganic Carriers for Bonded Phases... 58 2.2.10 Polymers as Stationary Phases ... 58 2.2.11 Porous Carbon ... 58 2.3 Retention with RP ... 58 2.3.1 Hydrophobic Properties of RP ... 59 2.3.2 RP with Shape Selectivity ...60 2.3.3 Influence of Analyte Structure on Retention ... 62 2.3.4 Influence of Eluent Composition ...64 2.3.5 Acetonitrile or Methanol ...64 2.3.6 Influence of Temperature ...68 2.3.7 Characterization and Comparison of RP ...69 References ... 74

in bioanalysis. All commonly used RP are based on silica. Their selectivity and efficiency depend on the physical and chemical properties of the base material silica and on the type and means of bonding of the alkyl groups for RP systems.

In literature there are many discussions on properties required for an optimal RP. According to Melander and Horvath [2], the ideal stationary phase for RP chromatography (RPC) should have the following properties:

A high efficiency and allows for rapid analysis.

Adequate stability and long life under widely varying operating conditions.

The column should permit the modulation of retention behavior over a wide range of

conditions. This means that the stationary phase is inert and does not exhibit specific inter- actions with certain functional groups of solutes with the concomitant advantage of rapid adsorption–desorption kinetics. Well-prepared hydrocarboneous-bonded phases should have properties that approach these requirements, which correspond to an ideal stationary phase.

The column material must be available with different mean pore diameters to allow effi-

cient separation of samples that fall in different molecular weight ranges.

The homogeneity of the surface should be such that the free-energy changes and the

adsorption–desorption rate constants associated with the chromatographic process on the molecular level fall within a narrow range.

The efficiency and the speed of analysis are related to the particle diameter of the stationary phase.

Nowadays, chromatographic columns are packed with stationary phases with average particle diameters below 10 μm. The tendency is to use even smaller particles, down to less than 2 μm.

The required theoretical plates for a given separation are then generated in much shorter columns, improving the speed of the analysis and the detection sensitivity. A discussion on the correlation of the particle diameter, the speed of the analysis, the column length and the required pressure drop is beyond the scope of this chapter and will be discussed elsewhere, e.g., in monographs on chro- matographic theory [3].

Other properties postulated for an optimal stationary phase depend on the carrier material—

almost exclusively silica—its physical and chemical properties, the type of bonded group (mainly octyl, C8 or octadecyl, C18), and the chemistry of the bonding reaction. There has been much improvement in our knowledge of the preparation of silica and the binding process in the course of RP development in the last 40 years approaching the desired goal. However, this is not a steady state and new and better, or at least different RPs, will become available. It is estimated that presently about more than 500 different RPs are on the market. Therefore, it will be beyond the scope of this chapter to summarize the commercially available RPs in a type of market overview. A discussion on the properties of RPs follows and an attempt is made to relate these properties to the type of silica carrier, the type of bonded organic moiety, and the means of binding reaction. Of course, this would not be possible without demonstrating and discussing the retention behavior of different classes of analytes in RPC.

The properties of RPs are determined by the Properties of the silica

Its specific surface area

Its pore size distribution

Its packing density

Its surface chemistry

Chemical modification

The type of silane

Its reactivity

The functional groups (alkyl; phenyl; fluoro; etc.)

The course of reaction leading to

Different carbon content

Surface coverage

Residual silanols concentration

End-capping or otherwise

2.2 the Base MaterIal: sIlICa

Different types of silica are used in the preparation of RPs. Silica are prepared by a condensation processes of soluble silicates like water glass, by destabilization of colloidal dispersions, or by polycondensation of ortho esters of silicic acid [4–6]. The formed hydrogels are aged, purified, and dehydrated to form xerogels. In the beginning (late 1960s) irregular silica prepared from sodium silicate was used (brand names, e.g., Lichrosorb, Bondapak, etc.). When these processes are per- formed in suspension between two immiscible liquids, spherical particles are obtained. The particle size can be adjusted by the stirring velocity. Spherical silica particles have also been prepared by emulsion polymerization in organic polymers. The organic part is removed by high temperatures, inducing a sintering process of initial nanospheres resulting in silica with much smaller porosities and higher packing densities (brand names, e.g., Hypersil, Spherisorb, Zorbax, etc.). The starting materials for these silica may contain heavy metals like iron, titanium, and alumina; consequently they have to be purified before use. As silica is a colloidal system, it changes its properties through weathering. To circumvent these problems, acid wash and rehydroxylation became popular in the 1980s. In the 1990s, very pure and metal-free silica were prepared, beginning with tetraethoxysi- lane (TEOS). These materials are optimal for RPC of basic solutes (brand names, e.g., Kromasil, Prontosil, Symmetry, etc.). By polycondensation of TEOS with alkylethoxysilanes, silica with an improved stability in alkaline solution can be obtained [7] (brand name, e.g., Xterra).

The physical properties of silica are determined by its specific surface area, pore volume, aver- age pore diameter, porosity, and the particle diameter and shape [8]. The latter two are responsible for the efficiency, the physical stability and the pressure drop of the packed columns and do not contribute to retention and selectivity.

2.2.1 pHysical paraMeters

Mostly fully porous particles are used in HPLC. The specific surface area and the pore diameter are interconnected in such a way that with an increasing surface area the average pore diameter decreases. Standard silica for HPLC have specific surface areas between 200 and 350 m2/g and aver- age pore diameters of 10–12 nm (100–120 Å). The surface area is exclusively within the pores. The outer surface of the spheres (geometrical surface) is negligible (<0.1%) for particles with diameters

>3 μm. For special applications, like in protein analysis, wide pore silica has to be applied with pore diameters from 30 up to 100 nm and corresponding surface areas between 100 and 10 m2/g. As wide pore materials are prepared from standard 10 nm materials by a hydrothermal process, these materials still contain a portion of smaller pores. The average pore size is given by a distribution often spanning one decade.

The pore volume of the silicas is in the range of 0.7–1.0 mL/g. In connecting these physical parameters, it becomes obvious that the wall between the pores is extremely thin. Calculations demonstrate [9] that the wall thickness (spread to a two-sided sheet) would be around 2.5 nm, or more realistically, as silica is formed by the condensation of nanoparticles, the wall is composed of spheres with a diameter of 8 nm. The model of plain cylindrical pores is used for calculations only and does not reflect a real picture.

The specific pore volume is important in size-exclusion chromatography (SEC) [10] because the separation takes place there. In retentive chromatography, it is necessary to provide the surface area

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