Laboratory assessment and diagnosis of congenital viral infections

Reproductive Toxicology 21 (2006) 350–382

Review

Laboratory assessment and diagnosis of congenital viral infections:Rubella, cytomegalovirus (CMV), varicella-zoster virus (VZV),

herpes simplex virus (HSV), parvovirus B19 andhuman immunodeficiency virus (HIV)

Ella Mendelson a,∗, Yair Aboudy b, Zahava Smetana c, Michal Tepperberg d, Zahava Grossman e

a Central Virology Laboratory, Ministry of Health and Faculty of Life Sciences, Bar-Ilan University, Chaim Sheba Medical Center, Tel-Hashomer, 52621, Israelb National Rubella, Measles and Mumps Center, Central Virology, Laboratory, Ministry of Health, Chaim Sheba Medical Center, Tel-Hashomer, Israel

c National Herpesvirus Center, Central Virology Laboratory, Ministry of Health, Chaim Sheba Medical Center, Tel-Hashomer, Israeld CMV Reference Laboratory, Central Virology Laboratory, Ministry of Health, Chaim Sheba Medical Center, Tel-Hashomer, Israel

e National HIV, EBV and Parvovirus B19 Reference Laboratory, Central Virology Laboratory, Ministry of Health,Chaim Sheba Medical Center, Tel-Hashomer, Israel

Received 14 October 2004; received in revised form 30 January 2006; accepted 7 February 2006

Abstract

Viral infections during pregnancy may cause fetal or neonatal damage. Clinical intervention, which is required for certain viral infections, relieson laboratory tests performed during pregnancy and at the neonatal stage. This review describes traditional and advanced laboratory approachesand testing methods used for assessment of the six most significant viral infections during pregnancy: rubella virus (RV), cytomegalovirus(CMV), varicella-zoster virus (VZV), herpes simplex virus (HSV), parvovirus B19 and human immunodeficiency virus (HIV). Interpretation ofthe laboratory tests results according to studies published in recent years is discussed.© 2006 Elsevier Inc. All rights reserved.

Keywords: Laboratory diagnosis; Congenital viral infections; Rubella; Cytomegalovirus (CMV); Varicella-zoster virus (VZV); Herpes simplex virus (HSV);Parvovirus B19; Human immunodeficiency virus (HIV)

Contents

1. General introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3522. Rubella virus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 352

2.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3522.1.1. The pathogen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3522.1.2. Immunity and protection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3522.1.3. Laboratory assessment of primary rubella infection in pregnancy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3552.1.4. Pre- and postnatal laboratory assessment of congenital rubella infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 357

2.2. Laboratory assays for assessment of rubella infection and immunity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3582.2.1. Rubella neutralization test (NT) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3582.2.2. Hemagglutination inhibition test (HI) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3582.2.3. Rubella specific ELISA IgG. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3582.2.4. Rubella specific ELISA IgM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3582.2.5. Rubella specific IgG-avidity assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3592.2.6. Rubella virus isolation in tissue culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359

∗ Corresponding author. Tel.: +972 3 530 2421; fax: +972 3 535 0436.E-mail address: [email protected] (E. Mendelson).

0890-6238/$ – see front matter © 2006 Elsevier Inc. All rights reserved.doi:10.1016/j.reprotox.2006.02.001

mailto:[email protected]

dx.doi.org/10.1016/j.reprotox.2006.02.001

E. Mendelson et al. / Reproductive Toxicology 21 (2006) 350–382 351

2.2.7. Rubella RT-PCR assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3592.3. Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359

3. Cytomegalovirus (CMV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3603.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 360

3.1.1. The pathogen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3603.1.2. Laboratory assessment of CMV infection in pregnant women. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3603.1.3. Prenatal assessment of congenital CMV infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 360

3.2. Laboratory assays for assessment of CMV infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3613.2.1. CMV IgM assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3613.2.2. CMV IgG assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3613.2.3. CMV IgG avidity assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3613.2.4. CMV neutralization assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3623.2.5. Virus isolation in tissue culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3623.2.6. Detection of CMV by PCR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3623.2.7. Quantitative PCR-based assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 362

3.3. Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3634. Varicella-zoster virus (VZV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363

4.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3634.1.1. The pathogen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3634.1.2. Assessment of VZV infection in pregnancy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3634.1.3. Prenatal and perinatal laboratory assessment of congenital VZV infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 364

4.2. Laboratory assays for assessment of VZV infection and immunity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3644.2.1. VZV IgG assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3644.2.2. VZV IgM assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3644.2.3. Virus detection in clinical specimens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3644.2.4. Virus isolation in tissue culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3644.2.5. Direct detection of VZV antigen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3644.2.6. Molecular methods for detection of viral DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 364

4.3. Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3655. Herpes simplex virus (HSV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365

5.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3655.1.1. The pathogen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3655.1.2. Laboratory assessment of HSV infection in pregnancy and in neonates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365

5.2. Laboratory assays for assessment of HSV infection and immune status . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3665.2.1. HSV IgG assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3665.2.2. HSV type-specific IgG assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3665.2.3. HSV IgM assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3665.2.4. Virus isolation in tissue culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3665.2.5. Direct antigen detection of HSV . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3665.2.6. Detection of HSV DNA by PCR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 366

5.3. Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3676. Parvovirus B19 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367

6.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3676.1.1. The pathogen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3676.1.2. Laboratory assessment of parvovirus B19 infection in pregnancy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3676.1.3. Prenatal laboratory assessment of congenital B19 infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367

6.2. Laboratory assays for assessment of parvovirus B19 infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3686.2.1. B19 IgM and IgG assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3686.2.2. Detection of viral DNA in maternal and fetal specimens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3686.2.3. Quantitative assays for detection of viral DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 369

6.3. Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3697. Human immunodeficiency virus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 369

7.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3697.1.1. The pathogen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3697.1.2. Importance of laboratory assessment of HIV infection in pregnancy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3707.1.3. Prenatal laboratory assessment of HIV infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 370

7.2. Laboratory assessment of HIV infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3707.2.1. HIV antibody assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3707.2.2. Detection of viral DNA in maternal and newborn specimens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 371

7.3. Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 372Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 372References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 372

352 E. Mendelson et al. / Reproductive Toxicology 21 (2006) 350–382

1. General introduction

Viral infections during pregnancy carry a risk for intrauterinetransmission which may result in fetal damage. The conse-quences of fetal infection depend on the virus type: for manycommon viral infections there is no risk for fetal damage, butsome viruses are teratogenic while others cause fetal or neonataldiseases ranging in severity from mild and transient symptomsto a fatal disease. In cases where infection during pregnancyprompts clinical decisions, laboratory diagnostic tests are anessential part of the clinical assessment process. This reviewdescribes the six most important viruses for which laboratoryassessement during pregnancy is required and experience hasbeen gained over many years. Rubella virus and CMV are ter-atogenic viruses, while VZV, HSV, parvovirus B19 and HIVcause fetal or neonatal transient or chronic disease.

The ability of viruses to cross the placenta, infect the fetusand cause damage depends, among other factors, on the mother’simmune status against the specific virus. In general, primaryinfections during pregnancy are substantially more damagingthan secondary infections or reactivations.

Laboratory testing of maternal immune status is required todiagnose infection and distinguish between primary and sec-ondary infections. Assessment of fetal damage and prognosisrequires prenatal laboratory testing primarily in those caseswhere a clinical decision such as drug treatment, pregnancy ter-m

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Since the algorithm for maternal and fetal assessment and theinterpretation of tests results vary from one virus to another, wehave described the approach to each viral infection in a separatechapter. Figs. 1–6 depict the most common algorithms used forthe laboratory diagnosis of each of the viral infections

2. Rubella virus

2.1. Introduction

2.1.1. The pathogenRubella is a highly transmissible childhood disease which

can cause large outbreaks every few years. It is a vaccine pre-ventable disease and in developed countries outbreaks are mostlyconfined to unvaccinated communities [1]. Rubella reinfectionfollowing natural infection is very rare. Rubella virus (RV) isclassified as a member of the togaviridae family and is the onlyvirus of the genus rubivirus [2]. Hemagglutinating activity and atleast three antibody neutralization domains were assigned to theearly proteins E1 and E2 [3–5]. At least one weak neutralizationdomain was identified on E2 [5].

The main route of postnatal virus transmission is by directcontact with nasopharyngial secretions [6]. Postnatal RV infec-tion is a generally mild and self-limited illness [6–8], but primaryRV infections during the first trimester of pregnancy have highteratogenic potential leading to severe consequences, known ascoi(

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ination or intrauterine IgG transfusion must be taken.This review describes basic virological facts and explains

he laboratory approaches and techniques used for the diagnos-ic process. It aims at familiarizing physicians with the rationalehind the laboratory requests for specific and timely specimensnd with the interpretation of the tests results including its lim-tations.

The laboratory methods used for assessment of viral infec-ions in general are of two categories: serology and virusetection. Serology is very sensitive but often cannot conclu-ively determine the time of infection, which may be criticalor risk assessment. Traditional serological tests, which mea-ure antibody levels without distinction between IgM and IgG,sually require two samples for determination of seroconver-ion or a substantial rise in titer. The modern tests can dis-inguish between IgG and IgM and may allow diagnosis inne serum sample. However, biological and technical difficul-ies are common and may cause false positive and false neg-tive results. The properties of all serological assays used forach of the viruses will be described in detail in the followinghapters.

Virus detection is used primarily for prenatal diagnosis. Inva-ive procedures must be used to obtain samples representing theetus such as amniotic fluid (AF), cord blood and chorionic villiCV). The traditional “gold standard” assay for virus detectionsed to be virus isolation in tissue culture, but other, more rapidnd sensitive methods were developed in recent years. Amonghe new methods are direct antigen detection by specific anti-odies and amplification and detection of viral nucleic acids.he general characteristics of all laboratory assays described in

his article are summarized in Table 1.

ongenital rubella syndrome (CRS) which may occur in 80–85%f cases [8,9]. It should be emphasized that more than 50% of RVnfections in non-immunized persons in the general populationand in pregnant women) are subclinical [6–9].

.1.2. Immunity and protectionAntibody level of 10–15 international units (IU) of IgG per

illilitre is considered protective. Naturally acquired rubellaenerally confers lifelong and usually high degree of immu-ity against the disease for the majority of individuals [10,11].ubella vaccination induces immunity that confers protection

rom viraemia in the vast majority of vaccinees, which usuallyersists for more than 16 years [10–12]. A small fraction of theaccinees fail to respond or develop low levels of detectablentibodies which may decline to undetectable levels within 5–8ears from vaccination [13–17].

Several methods are used to determine immunity (Table 1).eutralization test (NT) and hemagglutination inhibition test

HI) correlate well with protective immunity, but since they areifficult to perform and to standardize, they were replaced byhe more rapid, facile and sensitive enzyme-linked immunosor-ant assay (ELISA) [5,18]. In our experience (unpublished data),here is a clear distinction between antibody levels measuredsing ELISA, and antibody levels measured using functionalssays such as NT and HI. Moreover, standardization of antiV antibody assays using different techniques and a varietyf antigens (i.e., whole virus, synthetic peptides, recombinantntigen, etc.) has not been achieved, leading to uncertaintiesegarding the antibody levels that confer immunity and protec-ion against reinfections and against virus transmission to theetus [19–26]. Most of the reinfection cases (9 out of 18 cases;


Table 1Summary and characteristics of the laboratory tests used for assessment of viral infections in pregnancy

Laboratory test Test principles Clinicalsamples

Technical advantages Technicaldisadvantages

limitations Interpretation ofpositive results

SerologyNeutralization (NT) Inhibition of virus

growth in tissueculture by Aba

Maternalblood

Corresponds withprotection

Laborious, notvery sensitive, notAb class-specific

Done only inreference laboratories

Neutralizingantibodies are presentat a certain titer

Hemagglutinationinhibition (HI)

Prevention ofhemagglutination bybinding of Ab toviral Agb

Maternalblood

Accurate andcorresponds withprotection

Laborious, not Abclass specific

Used only for rubella,done only inspecialized labs

HI antibodies arepresent at a certaintiter

ELISA IgM Detection of virusspecific Ab bound toa solid phase by alabled secondaryanti-IgM Ab

Maternalblood, fetalblood,newborn blood

Fast and sensitive,commercialized,automated

None False positive andfalse negative

IgM antibodies arepresent

ELISA IgG Detection of virusspecific Ab bound toa solid phase by alabled secondaryanti-IgG Ab

Maternalblood,newborn blood


None None IgG antibodies arepresent (sometimeswith units)

IgG avidity(ELISA)

Removal of lowavidity IgG Abwhich results in areduced signal

Maternalblood


Not manyavailablecommercially

No interpretation forresults outside theinclusion or exclusioncriteria

Low avidity: recentinfection; mediumavidity: not known;high avidity: probablyold infection

Immunofluores-cence (IFA;IFAMA, etc.)

Detection of IgG orIgM Ab which bindsto a spot of virusinfected cells on aslide by a labledsecondary Ab

Maternalblood, fetalblood,newborn blood

Can yield titer; shorttime

Manual, reading issubjective

Unsuitable for testinglarge numbers

Antibodies are presentat a certain titer

Western blot (WB) Separated viralproteins attached toa nylon membranereact with patient’sserum and detectedby labledanti-human Ab

Maternalblood infant’sblood

Detects antibodyspecific to a viralprotein

Laborious Not very sensitive Antibodies specific tocertain viral antigensare present

Virus detectionVirus isolation in

tissue cultureInnoculation ofspecific tissuecultures withclinical samples andwatching for CPEc

Any clinicalsample whichmay containvirus

Detects and isolateslive virus

Very labouriousSlow

Insensitive, done onlyin virology labs

Live virus is presentin the clinical sample

Direct antigendetection

Detection of a viralantigen in cells froma clinical sample byIFA or ELISA

For IFA: cellsfrom clinicalsamples. ForELISA: anysample

Fast and simple Not sensitive Not sensitive, lowpositive predictivevalue

The sample mostlikely contains livevirus

Shell-vial assay Innoculation ofspecific tissuecultures withclinical samples,then fixation anddetection of viralcell-bound antigenby IFA


Detects live virus;rapid: results within16–72 h

Labourious;requires highskills; usesexpensivemonoclonal Abs

Not highly sensitive;done only in virologylabs

Live virus is presentin the clinical sample

Molecular assaysPCR; RT-PCR Enzymatic

amplification ofviral nucleic acidand detection ofamplified sequences


Fast, simple, can beautomated; verysensitive

Very prone tocontaminations

False positive bycontamination; maydetect latent virus

Viral nucleic acid ispresent in the sample,not known if live virusis present


Table 1 (Continued )

Laboratory test Test principles Clinicalsamples

Technical advantages Technicaldisadvantages

limitations Interpretation ofpositive results

Real-timePCR/RT-PCR

Detection ofaccumulating PCRproducts by afluorescent dye orprobe in aspecializedinstrument


Very fast, simple notprone tocontaminations; canbe quantitative

Expensiveinstruments

Sometimes toosensitive,interpretation of verylow resultquestionable

Viral nucleic acid ispresent in the sample(at a certain amount),not known if live virusis present

In situhybridization

Detection of viralnucleic acid insmears or tissuesections by labledprobes

Cells or tissuefrom clinicalsamples

Sensitive and specific Difficult toperform

Done only inspecializing labs


In situ PCR Detection of viralnucleic acid insmears or tissuesections by PCRusing labled primers

Cells or tissuefrom clinicalsamples

Sensitive and specific Difficult toperform

Doe only inspecializing labs


a Antibody.b Antigen.c Cytopathic effect.

50%) which were detected during an outbreak in Israel in 1992occurred in the presence of low neutralizing antibody titers of1:4 (cut off level), and sharp decline in the reinfection rate corre-lated with the presence of higher titers of neutralizing antibodies(unpublished data). Reinfection rates following vaccination are

considerably higher than following natural infection, rangingbetween 10% and 20% [19].

Many developed countries adopted the infant routine vacci-nation policy using MMR (mumps measles and rubella) vaccinedesigned to provide indirect protection of child-bearing age

F m shoa womw nth ob tion.o ed onp testsradvs

ig. 1. Algorithm for assessment of rubella infection in pregnancy: the algorithnd IgG. If the maternal blood is IgM negative the IgG result determines if theoman should be retested monthly for seroconversion till the end of the 5th moe an IgG avidity assay on the same blood sample to estimate the time of infecr recurrent infection. Medium AI is inconclusive and the test should be repeatositive and IgG negative, recent primary infection is suspected and the same
esults remain the same (IgM+ IgG−), then the IgM result is considered non-specificnd should be followed to the end of the 5th month as stated above). If the woman hasiagnosis should take place if the woman wishes to continue her pregnancy. Determinalue. Post natal diagnosis is based on the newborn’s serology (IgM for 6–12 m anecretions.
ws a stepwise procedure beginning with testing of the maternal blood for IgMan is seropositive (immune) or seronegative (not immune). If not immune thef pregnancy. If the maternal blood is IgM and IgG positive the next step wouldLow avidity index (AI) indicates recent infection while high AI indicates pasta second blood sample obtained 2–3 weeks later. If the maternal blood is IgM

should be repeated on a second blood sample obtained 2–3 weeks later. If the
, indicating that the woman has not been infected (however she is seronegativeseroconverted (IgM+ IgG+), recent primary infection is confirmed and prenatalation of IgM in cord blood is the preferred method with the highest prognostic
d IgG beyond age 6 m) and on virus isolation from the newborn’s respiratory


Fig. 2. Algorithm for assessment of CMV infection in pregnancy: the algorithm shows a stepwise procedure which begins with detection of IgM in maternal blood.If the maternal blood is IgG positive, an IgG avidity assay on the same blood sample should be performed to estimate the time of infection. Low avidity index (AI)indicates recent primary infection and prenatal diagnosis should follow. Medium or high AI is mostly inconclusive, especially if the maternal blood was obtained onthe second or third tremester. Continuation of the assessment is based on either maternal blood or fetal prenatal diagnosis. If the first maternal blood was IgM positivebut IgG negative, a second blood sample should be obtained 2–3 weeks later. If the IgG remains negative then the IgM is considered non-specific. If the womanhas seroconverted and developed IgG, primary infection is confirmed and prenatal diagnosis should follow. For prenatal diagnosis amniotic fluid (AF) should beobtained not earlier than the 21st week of gestation and 6 weeks following seroconversion. Fetal infection is assessed by virus isolation using standard tissue cultureor shell-vial assay, and/or by PCR detection of CMV DNA. Positive result by either one of these tests indicates fetal infection.

women regardless of vaccination status. However, in Israel andin other countries with high vaccination coverage, RV still cir-culates and may cause reinfections in vaccinated women whoseimmunity has waned [19,20,23, unpublished data].

2.1.3. Laboratory assessment of primary rubella infectionin pregnancy

Assessment of primary rubella infection in pregnant womenrelies primarily on the detection of specific maternal IgM anti-bodies in combination with either seroconversion or a >4-foldrise in rubella specific IgG antibody titer in paired serum sam-ples (acute/convalescent) as shown in Fig. 1. Today, due tothe high sensitivity of the ELISA-IgM assays low levels ofrubella specific IgM are detected more frequently, leading to

an increase in the number of therapeutic abortions and reducingthe number of CRS cases. However, frequently the low levelof IgM detected is not indicative of a recent primary infec-tion for several reasons: (a) IgM reactivity after vaccinationor primary rubella infection may sometimes persist for up toseveral years [27–29]; (b) heterotypic IgM antibody reactivitymay occur in patients recently infected with Epstein Barr virus(EBV), cytomegalovirus (CMV), human parvovirus B19 andother pathogens, leading to false positive rubella IgM results[30–35]; (c) false positive rubella specific IgM response mayoccur in patients with autoimmune diseases such as systemiclupus erythematosus (SLE) or juvenile rheumatoid arthritis, etc.,due to the presence of rheumatoid factor (RF) [36,37]; (d) lowlevel of specific rubella IgM may occur in pregnancy due to

F are shs early( al infd ht) shn

ig. 3. Algorithm for assessment of VZV infection in pregnancy: two situationserology (IgM and IgG in maternal blood) and by virus isolation or detection inpositive virus isolation/detection test and/or maternal seroconversion), then fetetection or PCR. (2) Exposure of a pregnant woman to a varicella case (top rigo IgG (not immune) she should receive VZIG within 96 h from exposure.

own: (1) clinical varicella in a pregnant woman (top left) should be assessed bydermal lesions. If either of those approaches confirms maternal VZV infection

ection can be assessed by virus detection in amniotic fluid using direct antigenould prompt maternal IgG testing within 96 h from exposure. If the mother has


Fig. 4. Algorithm for assessment of HSV infection in pregnancy and in neonates: the algorithm shows two complementary approaches to the confirmation of genitalHSV infection in pregnant women. (1) If genital lesions are present (top right), virus isolation and typing is the preferred diagnostic approach. A positive womanshould be examined during delivery for genital lesions. If normal delivery has taken place, the newborn should be examined for HSV infection symptoms and testedby virus isolation or PCR using swabs taken from skin, eye, nasopharynx and rectum or CSF. Detection of IgM in the newborn’s blood also confirms the diagnosis.Negative infants should be followed for 6 month. (2) Serology (top left) is a stepwise procedure beginning with maternal IgM and IgG testing. The interpretation ofthe results is shown: low positive or negative IgM in the presence of IgG indicates previous infection with the same virus type (reactivation), or recurrent infectionwith the other virus type. Type-specific serology may resolve the issue. If the IgG test is negative, in the presence or absence of IgM, a second serum sample shouldbe obtained to observe seroconversion. If the IgG remains negative then no infection occurred. If the woman seroconverted, type specific serology can identify theinfecting virus type.

Fig. 5. Algorithm for assessment of parvovirus B19 infection in pregnancy: the algorithm shows a stepwise procedure beginning with maternal serology followingclinical symptoms in the mother or in the fetus or maternal contact with a clinical case. Negative IgM and positive IgG indicate past infection, but if the IgG is highrecent infection cannot be ruled out. In all other cases a second serum sample should be obtained and retested. Only in the case of repeated negative results for bothIgG and IgM recent infection with B19 can be ruled out. In all other cases the fetus should be observed for clinical symptoms and if present tested for B19 infectionby nested PCR or rt-PCR performed on amniotic fluid or fetal blood. Positive result confirms fetal infection while negative result suggests that the fetus was notinfected with B19.


Fig. 6. Algorithm for assessment of HIV infection in pregnancy and in newborns to infected mothers: (1) diagnosis of maternal infection (top left) is by the routineprotocol (testing first by EIA and confirming by WB or IFA). If the mother is positive she should be treated as described in the text. (2) A newborn to an HIVpositive mother (top right) should be tested at birth by DNA PCR on a blood sample. The results, whether positive or negative, should be confirmed by retestingeither immediately (if positive) or 14–60 days later (if negative). If positive the newborn should be treated while if negative testing should be repeated at 3–6 monthsand again at 6–12 months. As a general rule, any PCR positive test in a newborn should be repeated on two different blood samples. After 12 months serologicalassessment can replace the PCR test as the infant has lost its maternal antibodies.

polyclonal B-cells activation trigerred by other viral infections[33,35,38].

False negative results may also occur in samples taken tooearly during the course of primary infection. Thus, the presenceor absence of rubella specific IgM in an asymptomatic patientshould be interpreted in accordance with other clinical and epi-demiological information available and prenatal diagnosis maybe required.

A novel assay developed recently to support maternal diag-nosis is the IgG avidity assay (Table 1) which can differentiatebetween antibodies with high or low avidity (or affinity) to theantigen. It is used when the mother has both IgM and IgG in thefirst serum collected (Fig. 1). Following postnatal primary infec-tion with rubella virus, the specific IgG avidity is initially lowand matures slowly over weeks and months [39–41]. Rubellaspecific IgG avidity measurement proved to be a useful toolfor the differentiation between recent primary rubella (clinicaland especially subclinical infection), reinfection, remote rubellainfection or persistent IgM reactivity. This distinction is crit-ical for the clinical management of the case, since infectionprompts a therapeutic abortion, reinfection requires fetal assess-ment, while remote infection or non-specific IgM reactivity carryno risk to the fetus [39–42].

2.1.4. Pre- and postnatal laboratory assessment ofcongenital rubella infection

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chorionic villi (CV) samples or amniotic fluid (AF) specimens.The laboratory methods used for virus detection are virus iso-lation in tissue culture or amplification of viral nucleic acids byRT/PCR (Table 1). However, using those methods for detectionof rubella virus in AF and CV might be unreliable, particularlyin AF samples due to low viral load. Studies showed that rubellavirus may be present in the placenta but not in the fetus, or itcan be present in the fetus but not in the placenta, leading tofalse negative results [6,43,44]. Thus, according to one opin-ion, detection of rubella virus in AF or CV does not justify therisk of fetal loss following these invasive procedures [45], whileaccording to another opinion, laboratory diagnosis of fetal infec-tion should combine a serological assay (detection of rubellaspecific IgM) with a molecular method (viral RNA detection) inorder to enhance the reliability of the diagnosis [46]. A recentstudy showed 83–95% sensitivity and 100% specifity for detec-tion of RV in AF by RT/PCR [47].

Postnatal diagnosis of congenital rubella infection [9,27,36]is based on one or more of the following:

a. Isolation of rubella virus from the infant’s respiratory secre-tions.

b. Demonstration of rubella specific IgM (or IgA) antibodies incord blood or in neonatal serum, which remain detectable for6–12 months of age.

c. Persistence of anti-rubella IgG antibodies in the infant’s

r

Maternal primary infection prompts testing for fetal infectionFig. 1). The preferred laboratory method for prenatal diagnosiss determination of IgM antibodies in fetal blood obtained byordocentesis [27,43]. Other options include virus detection in

serum beyond 3–6 months of age.

The principles, advantages and disadvantages of each labo-atory test, are described below.


2.2. Laboratory assays for assessment of rubella infectionand immunity

2.2.1. Rubella neutralization test (NT)Virus neutralization is defined as the loss of infectivity due to

reaction of a virus with specific antibody. Neutralization can beused to identify virus isolates or, as in the case of rubella diagno-sis, to measure the immune response to the virus [24,36,48]. Asa functional test, neutralization has proven to be highly sensitive,specific and reliable technique, but it can be performed only invirology laboratories which comprise only a small fraction ofthe laboratories performing rubella serology.

Rubella virus produces characteristic damage (cytopathiceffect, CPE) in the RK-13 cell line that was found most sen-sitive and suitable for use in rubella neutralization test. Othercells such as Vero and SIRC lines can be used if conditionsare carefully controlled [36]. Principally, 2-fold dilutions ofeach test serum are mixed and incubated with 100 infectiousunits of rubella virus under appropriate conditions. Then cellmonolayers are inoculated with each mixture and followed forCPE. Control sera possessing known high and low neutralizingantibody levels and titrations of the virus are included in eachtest run. The neutralization titer is taken as the reciprocal ofthe highest serum dilution showing complete inhibition of CPE[25,36].

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2.2.3. Rubella specific ELISA IgGThe ELISA technique was established for detection of an

increasing range of antibodies to viral antigens. In 1976, Volleret al. [58] developed an indirect assay for the detection of anti-viral antibodies. The technique has been successfully appliedfor the detection of rubella specific antibodies.

Almost all commercially available ELISA kits for the detec-tion of rubella specific IgG are of the indirect type, employingrubella antigen attached to a solid phase (microtiter polystyreneplates or plastic beads). The source of the antigen (peptide,recombinant or whole virus antigen) affects the sensitivity andspecificity of the assay. After washing and removal of unboundantigen, diluted test serum is added and incubated with theimmobilized antigen. The rubella specific antibodies present inthe serum bind to the antigen. Then, unbound antibodies areremoved by washing and an enzyme conjugated anti-human IgGis added and further incubation is carried out. The quantity of theconjugate that binds to each well is proportional to the concen-tration of the rubella specific antibodies present in the patient’sserum. The plates are then washed and substrate is added result-ing in color development. The enzymatic reaction is stoppedafter a short incubation period, and optical density (OD) is mea-sured by an ELISA-reader instrument. The test principle allowsthe detection of IgM as well by using an appropriate anti-humanIgM conjugate [53,57].

In most commercial ELISA IgG assays the results are auto-mta[rat

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.2.2. Hemagglutination inhibition test (HI)Until recently, assessment of rubella immunity and diagno-

is of rubella infection has been carried out mainly by the HIest which is based on the ability of rubella virus to agglutinateed blood cells [49]. HI test is labor intensive, and is currentlyerformed mainly by reference laboratories. HI is the “goldtandard” test against which almost all other rubella screeningnd diagnostic tests are measured. During the test, the agglu-ination is inhibited by binding of specific antibodies to theiral agglutinin. Titers are expressed as the highest dilutionnhibiting hemagglutination under standardized testing condi-ions [50–53].

The HI antibodies increase rapidly after RV infection sincehe test detects both, IgG and IgM class-specific antibodies.

titer of l:8 is commonly considered negative (cut off level::16) and a titer of ≥1:32 indicates an earlier RV infection oruccessful vaccination and immunity. Seroconversion is inter-reted as primary rubella infection, and a 4-fold increase in titeretween two serum samples (paired sera) in the same test series,s interpreted as a recent primary rubella infection or reinfec-ion [52]. Considerable experience has been accumulated overhe years in the interpretation of the clinical significance of HIiters [52–54], and the test results accurately correlate with clin-cal protection [5,18]. Although HI is generally considered asot sensitive enough, in certain situations it is still in use foresolution of diagnostic uncertainties.

Detection of rubella specific IgM class antibodies by HIest which requires tedious methods for purification of IgM oremoval of IgG [55,56], are no longer in use due to the develop-ent of a variety of rapid, easy to perform and sensitive methods,

f which ELISA is the most vastly used [18,57].

atically calculated and expressed quantitatively in interna-ional units (IU). When performed manually, the procedure takespproximately 3 h but automation has reduced it to about 30 min57–59]. It is important to note that in order to obtain reliableesults, determination of a significant change in specific IgGctivity in paired serum samples should always be performed inhe same test run and in the same test dilution.

The correlation between the ELISA and HI or NT titerss not always high. This may be explained by the fact thathe three methods detect antibodies directed to different anti-enic determinants [54]. Certain individuals fail to developntibodies directed to protective epitopes such as the neutral-zing domains of E1 and E2 due to a defect in their rubellapecific immune responses [21] but they do develop antibod-es directed to antigenic sub-regions of rubella virus proteins.LISA assays utilizing whole virus as antigen may fail to dis-

inguish between these different antibody specificities. Thus,eroconversion determined by ELISA based on a whole virusntigen does not necessarily correlate with protection againstnfection [52].

.2.4. Rubella specific ELISA IgMCommercially available ELISA kits for the detection of IgM

re mainly of two types:

. Indirect ELISA: The principle of the assay was describedabove for rubella IgG except for using enzyme labeled anti-human IgM as a conjugate. In this assay, false negative resultsmay occur due to a competition in the assay between specificIgG antibodies with high affinity (interfering IgG) while thespecific IgM have lower affinity for the antigen [31,32]. In the


new generation ELISA assays this is avoided by the additionof an absorbent reagent for the removal of IgG from the testserum. False positive results may occur if rheumatoid factor(RF: IgM anti-IgG antibodies) is present along with specificIgG in the test serum. Absorption or removal of RF and/orIgG is necessary prior to the assay to avoid such reactions[30–32,60].

b. IgM capture ELISA: In these assays anti-human IgM anti-body is attached to the solid phase for capture of serumIgM. Rubella virus antigen conjugated to enzyme-labeledanti-rubella virus antibody is added for detection. This typeof assay eliminates the need for sample pretreatment priorto the assay [32,61]. As for the rubella virus antigens, mostassays are based on whole virus extracts, but recent develop-ments led to production of recombinant and synthetic rubellavirus proteins [5,62].

2.2.5. Rubella specific IgG-avidity assayThis assay is based on the ELISA IgG technique and

applies the elution principle in which protein denaturant, mostlyurea (but also diethylamine, ammonium thiocyanate, guanidinehydrochloride, etc.) is added after binding of the patient’s serum.The denaturant disrupts hydrophobic bonds between antibodyand antigen, and thus, low avidity IgG antibodies produced dur-ing the early stage of infection are removed. This results in asignificant reduction in the IgG absorbance level [63]. The avid-i[

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Rubella virus can be grown in a variety of primary cells andcell lines [36,65], but RK-13 and Vero cell lines are the most sen-sitive and suitable for routine use. In these cell systems rubellavirus produces characteristic CPE. Since the CPE is not alwaysclear upon primary isolation, at least two successive subpas-sages are required [66]. When CPE is evident the identity of thevirus isolates should be confirmed using immunological or othermethods [36,65,67].

2.2.7. Rubella RT-PCR assayReverse transcription followed by PCR amplification (RT-

PCR) is a rapid, sensitive and specific technique for detectionof rubella virus RNA in clinical samples using primers fromthe envelope glycoprotein E1 open reading frame [45,46,68,69].Coding sequences for a major group of antigenic determinantsare located between nucleotides 731 and 854 of the E1 geneof RV strain M33. This region is highly conserved in variouswild type strains and is likely to be present in most clinicalsamples from rubella infected patients. Specific oligonucleotideprimers located in this region were designed for amplificationby RT-PCR [70–72]. Following rubella genomic RNA extractionfrom clinical specimens and RT-PCR amplification, the productis visualized by gel electrophoresis. Positive samples show aspecific band of the expected size compared to size markers[68,69,72].

A nested RT-PCR assay, in which the RT-PCR productfpstl

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ty index (AI) is calculated according to the following formula57]:

I = 100 × absorbance of avidity ELISA

absorbance of standard ELISA

The AI is a useful measure only when the IgG concentration inhe patient’s serum is not below 25 IU [39]. Low avidity (usuallyelow 50%) is associated with recent primary rubella infectionhile reinfection is typically associated with high avidity asresult of the stimulation of memory B cells (immunologicalemory) [39–41].In infants with CRS the low avidity IgG continues to be pro-

uced for much longer than in cases of postnatal primary rubella,here it lasts 4–6 week after exposure [39]. This may be used

or retrospective assessment of initially undiagnosed CRS cases.

.2.6. Rubella virus isolation in tissue cultureDiagnosis of prenatal or postnatal rubella infections are

ssentially based on the more reliable and rapid serologicalechniques. However, virus isolation is useful in confirming theiagnosis of CRS (Fig. 1) and rubella virus strain characteriza-ion required for epidemiological purposes. Rubella virus cane isolated using a variety of clinical specimens such as: res-iratory secretions (nasopharyngeal swabs), urine, heparinizedlood, CSF, cataract material, lens fluid, amniotic fluid, synovialuid and products of conception (fetal tissues: placenta, liver,kin, etc.) obtained following spontaneous or therapeutic abor-ion [6,36,44,64]. In order to avoid virus inactivation, specimenshould be inoculated into cell culture immediately or stored at◦C for not more than 2 days, or kept frozen (−70◦ C) for longereriods [36].

rom the first amplification reaction is re-amplified by internalrimers, was developed and shown to provide a higher level ofensitivity for the detection of rubella virus RNA [72]. However,he risk of contamination is markedly increased. The detectionimit of the RT-PCR assay is approximately two RNA copies.

Clinical specimens for rubella virus genome detectionnclude: products of conception (POC), CV, lens aspirate/biopsy,F, fetal blood, pharyngeal swabs and spinal fluid (CSF) orrain biopsy when the central nervous system (CNS) is involved68,69,73–75]. An additional advantage of RT-PCR is that it doesot require infectious virus [74]. RV is extremely thermo-labilend frequently is inactivated during sample transporation to theaboratory.

Finally, it should be noted that clinical samples may con-ain PCR inhibitors (such as heparin and hemoglobin), andhe extraction procedure itself may cause enzyme inhibition72,76,77]. This underscores the need and importance for strictnternal quality control during each step of the RT-PCR proce-ure and participation in external quality assessment programss of a high value.

.3. Summary

Rubella infection during pregnancy, although rare in coun-ries with routine vaccination programs, is still a problem requir-ng careful laboratory assessment. The laboratory testing shouldonfirm or rule-out recent rubella infection in pregnant womennd identify congenital rubella infections in the fetus or neonate.aternal infection is currently assessed by serological assays,

rimarily by ELISA IgM and IgG. Borderline results for the IgGssay can be further assessed by the HI or NT assays available


in reference laboratories. Confirmation of recent infection canbe sought using the IgG-avidity assay in addition to the othertests.

Intra-uterine infection is assessed by IgM assays in fetal bloodwhich can be accompanied by virus detection in CV or AF spec-imens, or, in case of induced abortion, in fetal tissue. Laboratoryassessment of congenital rubella infection in neonates relies onvirus detection by culture or RT-PCR in various clinical sam-ples taken early after birth, and by demonstration of IgM andlong-lasting IgG in neonatal serum. Due to the complexity ofthe current laboratory assays, cooperation between the physi-cian and the laboratory is of utmost importance to achieve areliable diagnosis.

An algorithm describing the laboratory diagnosis process forRV is shown in Fig. 1.

3. Cytomegalovirus (CMV)

3.1. Introduction

3.1.1. The pathogenCMV is a common pathogen which can cause primary and

secondary infections. CMV is a member of the herpesvirus fam-ily possessing a 235 kb double stranded linear DNA genome, acapsid and a loose envelope. Membranal glycoproteins embed-ded in the envelope carry neutralization epitopes. CMV canieferihait[t

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3.1.2. Laboratory assessment of CMV infection in pregnantwomen

CMV was recognized as the cause of fetal stillbirth followinga cytomegalic inclusion disease (CID) in the mid 1950s when itwas first grown in tissue cultures in three laboratories [80–82].Since then demonstration of CMV infection of the mother orfetus by laboratory testing has become an essential part of theassessment of pregnancies at risk [76,83]. Assessment of con-genital CMV infection begins with maternal serology whichshould establish recent primary or secondary infection (Fig. 2).

Not all maternal infections result in fetal transmission anddamage. Only 35–50% of maternal primary infections and0.2–2% of secondary infections lead to fetal infection, out ofwhich only 5–15% in primary infection and about 1% in sec-ondary infections are clinically affected [84–87]. Therefore,following maternal diagnosis, and if early pregnancy termina-tion was not chosen, subsequent prenatal diagnosis should takeplace using methods for virus detection in AF samples.

Demonstration of maternal infection relies on ELISA IgMand IgG assays and on CMV IgG avidity assay (Fig. 2). UnlikeHI and NT for rubella, for CMV there are currently no serolog-ical “gold standard” assays which can be used for confirmationand reassurance. Recently an attempt to find association betweenviral load in maternal blood and the risk for fetal infection didnot yield positive results [88].

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nfect all age groups usually causing mild and self-limited dis-ase. Its sero-prevalence in women of child-bearing age variesrom 50% to over 80%, with inverse correlation to socio-conomic levels. Primary CMV infection during pregnancy car-ies a high risk of intrauterine transmission which may resultn severe fetal damage, including growth retardation, jaundice,epatosplenomegaly and CNS abnormalities. Those who aresymptomatic at birth may develop hearing defects or learn-ng disabilities later in life. It is now recognized that intrauterineransmission may occur in the presence of maternal immunity78]. Pre-conceptional primary infection carries a high risk iden-ical to the risk of infection during early gestational weeks [79].

CMV, like other members of the herpesvirus family, estab-ishes a latent infection with occasional reactivations as well asecurrent infections in spite of the presence of immunity. How-ver, reactivation or recurrent infections carry a much lower riskor fetal infection and damage is much lower in such events.

The infectious cycle in vitro takes 24–48 h while in vivo thencubation period for postnatal infection can last for 4–8 weeks.he incubation period for congenital infection is not known and

he gestational age of congenital infection is currently definedy the maternal seroconversion, if known, which does not nec-ssarily reflect the actual timing of the fetal infection.

The host defense against CMV infection in immune-ompetent individuals combines cellular and humoral immuneesponses which together prevent a severe CMV disease in theast majority of infections. Antibodies of the IgM class areroduced immediately after primary infection and may last foreveral months. IgM can be produced in secondary infectionsn some cases. Antibodies of the IgG class are also producedmmediately after infection and last for life.

.1.3. Prenatal assessment of congenital CMV infectionMaternal infection during pregnancy prompts testing for fetal

nfection as outlined in Fig. 2. Prenatal CMV diagnosis can-ot rely on detection of fetal IgM since frequently the fetusoes not develop IgM [76,89–94]. On the other hand, becauseMV is excreted in the urine of the infected fetus, detection ofirus in the AF has proven to be a highly sensitive and reliableethod. Numerous studies have focused on the most appropriate

iming for performing amniocentesis which will yield the bestensitivity for detection of fetal infection [76,83,97–99]. Thesetudies clearly indicated that amniotic fluid should be collectedn 21–23 gestational week and at least 6–9 weeks past maternalnfection. If these requirements are met then the sensitivity ofetection of intrauterine infection can reach over 95% while theeneral sensitivity is only 70–80%. One study measured the sen-itivity for AF obtained at gestational weeks 14–20 and reportednly 45% [100]. Most of the studies state that the timing of themniocentesis is more critical for sensitivity than the laboratoryethods used to detect the virus in the AF.Initially, virus isolation in tissue culture and its more sophisti-

ated variation “Shell-Vial” technique (Table 1) were the leadingaboratory methods for detection of CMV in amniotic fluid.owever, during the late 1980s highly sensitive molecular meth-ds were developed for detection of specific viral DNA inlinical specimens such as dot-blot hybridization [101–103].hese methods were much faster, less laborious and repeat-ble compared to virus culturing. Performance of the biologicalnd molecular techniques in parallel assured that the preciousmniotic fluid sample will not be wasted and that false negativeesults will not be obtained by a technical problem in any ofhese “home-made” assays.


Since the early 1990s the polymerase chain reaction (PCR)has become the preferred method for CMV detection in amnioticfluid [95,96,104–106]. Problems with molecular contaminationleading to false positive results and the need to address prog-nostic issues, led finally to the development of quantitative PCRassays with the highly advanced real-time PCR (rt-PCR) as themost updated method (Table 1). Current studies deal with thecorrelation between the “viral load” in the amniotic fluid andthe pregnancy outcome, in an attempt to establish the prognos-tic parameters of this powerful technique.

The laboratory methods used for assessment of maternal andfetal CMV infection are described in detail below.

3.2. Laboratory assays for assessment of CMV infection

3.2.1. CMV IgM assaysIgM detection is a hallmark of primary infection although it

may also be associated with secondary infections [90,107–109].Major efforts were put into developing sensitive and reliableassays for IgM detection using ELISA. The technical and bio-logical obstacles and their solutions which were described forrubella IgM assays apply for CMV as well, including long-termpersistence of IgM antibodies [110–114].

The source of the viral antigen affects sensitivity and speci-ficity [113,115–122], but in the absence of a gold standard assay,comparisons between various commercially available assayswbaoalacmdba[

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since a “significant increase” is rarely defined for commercialELISA assays, it is up to the laboratory to define it.

3.2.3. CMV IgG avidity assaysThe IgG avidity assay was developed to circumvent diagnos-

tic problems as described for rubella [128–130]. It is performedwhen both IgM and IgG are positive on initial testing, but cannotbe performed on sera with very low IgG titers. Various commer-cial assays are calibrated in different ways for determination ofthe diagnostic threshold: some assays exclude recent infectionif the AI reaches a certain threshold level, yet others approverecent infection if the AI is lower than a certain threshold level.However, none of these assays can exactly determine when theinfection occurred or give any interpretation of results fallingoutside of its exclusion or inclusion criteria. Numerous studiespublished in recent years aimed at evaluating IgG-avidity assays(by commercial kits or “in-house” methods) for their ability toidentify or exclude recent primary CMV infection, and to predictcongenital infection. Concordance between different commer-cial assays for determination of low avidity was high (98–100%),but not for determination of high avidity (70%). Because the useof this assay is relatively new, some of these studies are describedin detail below.

One set of studies evaluated the ability of the assay to assessthe risk for fetal infection. [127,131–133]. In a cohort of womenconsidered at risk for transmitting CMV to their fetuses basedosttaa(cia

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ere based on multi-variant analyses of “consensus” resultsetween several assays. These studies demonstrated high vari-bility in specificity and sensitivity among assays and a high ratef discordance [123–126]. Thus, testing for IgM, particularly insymptomatic pregnant women, may frequently create a prob-em rather that solving it: borderline results or conflicting resultsmong two or more commercial kits are interpreted as incon-lusive and require further testing as described below. Otherethods, such as immunoblotting and IF assays (Table 1) were

eveloped to confirm positive IgM results and to distinguishetween specific and non-specific reactions [88]. However, thesessays did not gain vast usage because of lowered sensitivity127] and the lack of automation.

.2.2. CMV IgG assaysIgG assays which are currently based on ELISA, are generally

sed for determination of immune status but, unlike rubella,here is neither definition of a CMV-IgG international unit (IU)or of the protective antibody level. IgG assays may also helpo establish diagnosis of current CMV infection in suspectedecondary infections, or when the IgM result is inconclusive,y demonstration of IgG seroconversion or a significant IgGise between paired sera taken 2–3 weeks apart. This ability isimited in cases when women initially present with a high titer ofgG or when it is impossible or too late to obtain a second serumample. Commercial ELISA IgG assays are relatively simple,orrelate well with each other and most of them are quantitativeut are not yet internationally standardized. Commercial kits userbitrary units (AU) which differ from one assay to another andhus, to demonstrate an increase in antibody level, it is criticalo run the two samples in parallel in the same test. Additionally,

n demonstration of IgM or seroconversion, low avidity wastrongly associated with fetal infection (100% sensitivity) ifhe serum sample tested was collected at 6–18 weeks gesta-ion. Moderate or high AI levels were associated with 33%nd 11%, respectively, of cases with CMV genome-positivemniotic fluid, but with no fetal infection. Lowered sensitivity60–63%) for detection of primary infection was found for seraollected at 21–23 weeks gestation, since some of the mothers,nfected early in pregnancy, already developed moderate or highvidity.

Another set of studies [134–136] examined the ability of thegG-avidity assay to exclude those with past infection and there-ore with low risk of fetal transmission. Women with positiver equivocal IgM but without documented seroconversion wereested. High avidity was interpreted as remote infection whichid not occur within the last 3 months. The results of this seriesf studies also showed that congenital infection was stronglyssociated with low avidity, while moderate or high avidity weressociated with uninfected fetus. Additional studies further con-rmed the strong association between low avidity and primary

nfection, and thus risk for fetal infection, and between highvidity and past infection [130,136–140]. One study showedhe lack of full concordance between different commercial IgGvidity assays [141]. It showed that the ability of a commercialit to exclude recent infection by high avidity was restricted toI of >80% and to determine recent infection to AI of <20%.ny result in between those limits was inconclusive since seraith AI of 50–80% included 48 out of 257 (18%) women withhistory of past infection and 3 sera from 2 patients with a his-

ory of recent infection. Testing the latter three samples with aifferent kit yielded low avidity (30%).


In conclusion, the IgG-avidity assay is a powerful tool butit should be used and interpreted properly. The associationbetween low AI and recent primary infection with a high riskfor congenital infection is stronger than the association betweenmoderate or high AI and past infection with low risk. Inter-pretable results can be achieved mainly for sera obtained withinthe first 3–4 month of pregnancy. However both the inclusionand the exclusion approaches can be used and the IgG avidityassay is now implemented in a testing algorithm following theIgG test [142] as shown in Fig. 2.

3.2.4. CMV neutralization assaysNeutralizing antibodies appear only 13–15 weeks following

primary infection, thus the presence of high-titer of neutralizingantibodies during acute infection indicates a secondary ratherthan a primary infection. The neutralization assays have notreached a wide use as they are labor intensive, very slow andcannot be commercialized. Therefore, they are rarely performedby specialized reference laboratories. Attempts to correlate neu-tralization with specific response to the viral glycoprotein gBshowed promising results and were also commercialized in anELISA format [130,143–146]. However the utility of this assayrequires further studies.

3.2.5. Virus isolation in tissue cultureVirus is cultured from AF samples to assess fetal infection

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except for rare cases in which the monoclonal antibody does notrecognize the viral antigen [147–149].

Today, virus isolation from AF remains a key method fordemonstration of fetal infection and has been described exten-sively in many studies either exclusively or in conjunction withmolecular methods, particularly PCR [97,99,132,150–155]. Themain subject under investigation in recent years has been thecomparative sensitivity and specificity of the PCR and the virusisolation methods.

3.2.6. Detection of CMV by PCRDetection of viral DNA in clinical samples involves DNA

extraction and analysis. PCR has become the preferred methodfor rapid viral diagnosis in recent years. Its main disadvan-tage is the possible contamination leading to false positiveresults.

The PCR assay includes several components which can varyfrom test to test. Viral DNA can be extracted using in-housemethods or various commercial kits. The primers used can bederived from different viral genomic sites and the reaction con-ditions can be altered. For CMV, most assays utilize the early (E)or immediate-early (IE) genes which are highly conserved com-pared to the structural matrix or glycoprotein genes presentinghigher variability among wild-type isolates. To increase sensitiv-ity [95,156] some assays include a second round of amplificationusing nested or hemi-nested primers. The nested PCR is how-er

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Fig. 2). Since CMV is a very labile virus, samples for culturinghould be kept at 4–8 ◦C and transported to the laboratory within8 h to be tested immediately. Freezing AF at −20 ◦C destroysirus infectivity and freezing at or below −70 ◦C requires stabi-ization by 0.4 M sucrose phosphate [147].

Culturing CMV from AF which was stored and transportedppropriately, has always been considered the gold standardethod for detection of fetal infection having 100% speci-city. CMV can be isolated in human diploid fibroblast cellsither primary, such as human embryonic cells and human fore-kin cells, or continuous cultures such as MRC-5 and WI-38ells [147]. The diploid cells should be used at low passageumber to avoid loss of sensitivity. Tissue culture monolay-rs inoculated with the clinical samples should be maintainedor up to 6 weeks since CMV is a slow growing virus. Dur-ng this period blind passages should be performed using cellsather than culture supernatant since the virus is cell-bound.MV produces a typical CPE which can appear within 2–3ays and up to 6 weeks, depending on the virus concentra-ion in the clinical sample. The CPE is easily recognizable,ut IF assay using specific antibodies can confirm the presencef CMV.

An alternative, much shortened procedure called the “shell-ial assay” was developed in which inoculated cultures are spunown at low velocity for 40–60 min before incubation at 37 ◦C.his procedure enhances and speeds-up viral infection of theultured cells. Infected cells are then detected within 16–72 hy IF using monoclonal antibodies directed against early viralroteins synthesized shortly after infection. This method gainedide acceptance and is now used by most laboratories. Its sen-

itivity and specificity are highly comparable to virus isolation

ver more prone to molecular contamination and false-positiveesults.

The comparative specificity and sensitivity of PCR and virussolation is dependent upon technical parameters which varyrom one laboratory to another with relation to the overall med-cal set-up in which they are placed and the technical skills ofhe laboratory personnel. However, it is generally agreed that forMV, PCR is more sensitive than tissue culture isolation. PCR

s also a repeatable assay which is of great advantage in contro-ersial cases. Original samples kept frozen at or below −70 ◦Can be re-processed and extracted DNA can be re-tested or sento another laboratory for confirmation.

.2.7. Quantitative PCR-based assaysMany previous studies have shown that detection of CMV

NA in AF by itself does not predict the outcome of fetal infec-ion. Clinical measures such as ultrasonographic examinationsre a key component in fetal assessment, but might also fail toetect affected fetuses. In an attempt to address prognostic issuest was suggested that symptomatic fetuses can be distinguishedrom asymptomatic ones based on the viral load in the amnioticuid. Quantitative PCR methods were developed as “in-house”ssays or are available as commercial kits using various tech-ologies. The most up-to date technology is the real-time PCRssay in which the amplified sequences are detected by a fluo-escent probe in a real-time and quantitative manner [157–159].hese assays, performed by dedicated instruments, carry thedvantages of high sensitivity and specificity conferred by theybridization probe, and the lack of contamination by amplifi-ation products, since the reaction tubes are never opened aftermplification.


Few recent publications have addressed the prognostic valueof determination of viral load in AF with controversial results.Three studies [160–162] found no statistically significant differ-ence in viral load between symptomatic and asymptomatic fetalinfections. Yet other two studies reported predictive values forviral load [163,164]. In one of these two studies [164] the pres-ence of 103 or more CMV genome-equivalents per millilitres(GE/ml) predicted mother to child transmission with 100% prob-ability, and 105 GE/ml or more predicted symptomatic infection.In the second report [163] CMV DNA load with median of2.8 × 105 GE/ml was associated with major ultrasound abnor-malities while median values of 8 × 103 GE/ml was associatedwith normal ultrasound and asymptomatic newborn. The slightdiscordance between the two studies calls for further evaluationson a larger scale and underscores the need for standardization,since the quantitative assays may vary by orders of magni-tude using different methods or primers derived from differentgenomic regions [165,166].

3.3. Summary

Laboratory testing for determination of intrauterine CMVinfection involves several steps. Maternal primary or recurrentinfection is assessed by serology using IgM, IgG and IgG-avidityassays. In controversial cases a second blood sample should besought to demonstrate antibody kinetics typical of current infec-tmwgtQd

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and the fetus. Complications and sequelae include pneumo-nia, increased rate of prematurity abortions, congenital varicellasyndrome (CVS), neonatal varicella and herpes zoster duringthe first year of life [171–177]. Rarely VZV may cause a lifethreatening CNS infection. The risk of adverse effects for themother is greatest in the third trimester of pregnancy, while forthe fetus the risk is greatest in the first and second trimesters.The risk of CVS for all pregnancies continuing for 20 weeks isabout 1%, but is lower (0.4%) between weeks 0 and 12 and ishigher (2%) between weeks 13 and 20. Maternal infection after20 weeks and up to 36 weeks is not associated with adversefetal effect, but may present as shingles in the first few yearsof infant’s life indicating reactivation of the virus after a pri-mary infection. If maternal infection occurs 1–4 weeks beforedelivery, up to 50% of the newborns are infected and 23%of them develop clinical varicella. Severe varicella occurs ifthe infant is born within seven days of the onset of maternaldisease.

4.1.2. Assessment of VZV infection in pregnancyLaboratory diagnosis of VZV in pregnancy is required in two

situations: (a) the pregnant woman has developed clinical symp-toms compatible with chickenpox or herpes zoster (Shingles) (b)The pregnant woman was exposed to a chickenpox or a zostercase (Fig. 3).

If the pregnant woman has developed clinical symptomstsm

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ion and not of remote infection or a non-specific reaction. Ifaternal primary infection was established and the pregnancyas not terminated, prenatal diagnosis follows at 21–23 weeksestation and 6–9 weeks after seroconversion (if known). Detec-ion of CMV in AF is done by virus culturing and/or PCR.uantitative PCR is still not established for assessment of fetalamage and prognosis.

During the diagnostic process, which may last for severaleeks, collaboration between the laboratory and the physician

s of utmost importance. Appropriate timing of sampling, sam-le treatment, usage of validated assays under quality assessmentonditions, and correct interpretation of the results are all essen-ial for obtaining a reliable diagnosis.

The algorithm describing the laboratory diagnostic processor CMV is shown in Fig. 2.

. Varicella-zoster virus (VZV)

.1. Introduction

.1.1. The pathogenVaricella-zoster virus (VZV) is a common pathogen belong-

ng to the herpesvirus family which can establish latent infec-ions and subsequent reactivations. Primary infection, chicken-ox, is a common childhood disease. Reactivation is manifesteds zoster and occurs in the presence of anti VZV antibodies.pproximately 90% of the adult population is positive for VZV

ntibodies and studies on pregnant and parturient women foundetween 80% and 91% seropositivity [167–170].

Primary infection with VZV (chickenpox) during pregnancyarries a risk for clinical complications for both the mother

he infection should be confirmed by laboratory testing usingerology or virus detection by culturing, antigen detection orolecular methods.Assessment of VZV IgM which remains in the blood for 4–5

eeks is diagnostic. However, false positive results are com-on in the presence of high VZV IgG antibodies and virus

eactivations may also induce IgM. Therefore, determinationf IgG seroconversion or a 4-fold rise in VZV IgG titer shouldccompany the IgM test. Virus isolation or PCR from dermalesions can be attempted and, if positive, confirm the diagnosisFig. 3).

If the pregnant woman has reported exposure to a case ofhickenpox or zoster, prompt assessment of her immune statusy testing for IgG within 96 h from exposure should be doneince varicella-zoster immunoglobulin (VZIG) given as a pro-hylactic measure at the time of exposure is known to preventr reduce the severity of chickenpox [178,179].

Serological screening for IgG of women with negative orncertain histories of illness, who are planning a pregnancy,r of women who give history of recent contact with chicken-ox, has been suggested as a strategy for preventing CVS andeonatal VZV [180,181]. In a study conducted in our labora-ory 52 pregnant women were assessed for immunity to VZVollowing exposure to chickenpox and 25% of them were foundusceptible. The attack rate among the susceptible women was5% [181]. In another study, Linder et al. [182] reported thatn 327 pregnant women assessed for VZV immunity, 95.8% ofhe women who recalled chickenpox in themselves and 100% ofomen who recalled chickenpox in their children were seropos-

tive, and only 6.8% of the women with a lack or uncertainistory of exposure were seronegative. The screening strategy


might gain momentum due to the availability of VZV vaccine,as seronegative women can be vaccinated.

4.1.3. Prenatal and perinatal laboratory assessment ofcongenital VZV infection

If VZV infection of the mother during the first or the sec-ond trimester has been confirmed, the need to diagnose the fetusarises. Unfortunately, laboratory methods for fetal assessmentare of limited value. Assessment of fetal infection by determi-nation of VZV IgM in fetal blood is not widely performed. IgMmay be manifested in the fetus only after 24 weeks of gesta-tion, thus in case of intrauterine infection during early gestation,functional immunity may not be present in the fetus [183].Alternatively, determination of fetal infection can be done bydemonstration of VZV DNA in AF using PCR, but its presenceis not synonymous with development of CVS [183]. Only onein 12 infected fetuses will develop pathological signs, so inter-pretation of a positive VZV DNA result is problematic if thefetus appears normal upon ultrasonograpic examination [180].Diagnosis of clinical varicella in neonates is based on serology(IgM) and virus isolation or detection in vesicle fluid or in CSF(in case of CNS infection).

4.2. Laboratory assays for assessment of VZV infection andimmunity

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4.2.4. Virus isolation in tissue cultureVZV isolation in tissue culture is not a very sensitive assay

and the isolation of the virus from skin lesions is possible onlyfrom early vesicles. VZV is a very labile virus which growsslowly in selected tissue cultures. Specimens for cell culture(vesicle fluid, skin-lesion swabs and AF) should be transportedto the laboratory as soon as possible after collection [196–198].Swabs should be put in a vial containing virus transport mediumand AF should be placed in a sterile container. Both should bekept at 4–8 ◦C during transportation. Inappropriate conditionsduring transportation may easily reduce virus viability [199].

Upon receipt, the specimen is inoculated into semi-continuous diploid cells such as human diploid fibroblasts(MRC-5) and continuous cell lines derived from tumors ofhuman or animal tissue such as A549. CPE may appear within 2weeks. AF should be sampled after 18 weeks of pregnancy andafter complete healing of skin lesions of the mother. Confirma-tion of virus isolation can be done by immunoflurescent stainingusing monoclonal anti-VZV antibody. Shell vial cultures asdescribed for CMV improve the sensitivity of VZV detectionand allow rapid identification of positive samples within 1–3days [196,200].

4.2.5. Direct detection of VZV antigenImmunofluorescence or immunoperoxidase assays use mon-

oclonal or polyclonal antibodies directed to VZV antigens todfrahoan

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.2.1. VZV IgG assaysSeveral methods were applied for determination of VZV

ntibodies. In the past, specific IgG antibodies were measuredy the complement fixation (CF) assay, which can be usednly for diagnosis of recent infection or by the latex agglu-ination assay [184–188]. Today, highly sensitive and specificLISA and immunofluorescence (IF) methods are used foretermitation of VZV IgG antibodies. Several versions of theF assay exist (Table 1): fluorescent antibody to membrane anti-en (FAMA) and indirect fluorescent antibody to membranentigen (IFAMA) detect binding of antibodies to membranentigens in fixed VZV infected cells, and are highly specificnd sensitive [169,181,186,189]. ELISA is comparable to IFn sensitivity, specificity and cost and can be used for generalcreening purposes. Commercial ELISA kits for VZV IgG areenerally highly specific but demonstrate some false positiveesults compared to FAMA or IFAMA.

.2.2. VZV IgM assaysBoth the IF and ELISA methods are used for determination

f VZV IgM in the same manner as for IgG, except that theonjugate is an anti-human IgM rather than anti-human IgG.ommercial ELISA IgM kits may give false positive results asas described for RV and CMV.

.2.3. Virus detection in clinical specimensVirological methods for the diagnosis of VZV infection

nclude detection of infectious VZV, viral antigens and viralNA in clinical specimens. These include vesicular fluid, swabsr smears, AF in pregnant women or cerebrospinal fluid (CSF)n encephalitis cases [183,190–195].

etect VZV infection in epithelial cells isolated from AF, orrom suspected lesions (Table 1). This simple method allowsapid diagnosis (approximately 2 h) of VZV infection and isvailable to most laboratories [201,202]. It is highly specific andas sensitivity ranging from 73.6% to 86%. The preferred mon-clonal antibodies are those directed against the cell-membranessociated viral antigens [203]. Stained smears, in which multi-ucleated giant cells are seen are less sensitive, 60–75% [204].

.2.6. Molecular methods for detection of viral DNAPCR and hybridization methods to detect VZV DNA

equences are very sensitive and specific [183,191,205–208].odifications of the basic PCR technique have been used to

ncrease the sensitivity by using nested PCR assays. However,his assay is highly susceptible to contamination, leading to falseositive results. Rapid laboratory diagnosis is important whenhe CNS is involved, especially in cases with clinically confusedermal manifestations, and is crucial in neonates to prevent lethalutcome of disease. In recent years real-time PCR assays wereeveloped for VZV as for CMV and other viruses. Real-timeCR assays appear to have equal sensitivity as VZV nested PCRssays but are faster, easier and have markedly reduced risk forolecular contamination. Because of its high sensitivity com-

ared to other methods, PCR has become the most appropriateethod for detection of VZV DNA in AF and other specimens

183,206,209,210]. However, standardization is a major prob-em and should be done in order to avoid false positive or falseegative results.

In a study conducted in our laboratory AF samples werenalyzed either by PCR or real-time PCR in parallel to tissueulture isolation. The DNA amplification target was located in


the viral UL gene. VZV DNA was detected in the amniotic fluidof two (7.4%) out of 27 women who developed chickenpox inthe second trimester of pregnancy. One gave birth to a normalchild while no follow-up was available for the second woman.All amniotic fluid cell cultures were negative. No case of CVSoccurred following negative PCR results (unpublished data).

4.3. Summary

VZV infection during pregnancy poses a risk to the motherand fetus. Laboratory assessment of maternal infection is basedon serology (both IgG and IgM), and on virus detection inskin lesions. Exposure of a pregnant woman to a varicella caseprompts immediate assessment of her immune status to deter-mine the need for VZIG administration. The availability of aVZV vaccine may encourage the screening of women for VZVimmunity before conception.

There are no fully reliable methods to assess fetal infectionand damage. Fetal IgM and virus detection in AF are used, butfalse negative results are common and positive results do notnecessarily correlate with fetal damage.

An algorithm describing the laboratory diagnostic assays forVZV infection in pregnancy is shown in Fig. 3.

5. Herpes simplex virus (HSV)

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infections are associated with prematurity and fetal growthretardation.

Herpes simplex is a devastating infection in the neonate,with primary asymptomatic and symptomatic maternal infec-tion near delivery carrying a greater risk to the newborn thanrecurrent infections [218,219]. Fourty percent of women whoacquired genital HSV during pregnancy but did not completetheir seroconversion prior to the time of delivery will infecttheir newborns. Transmission of genital herpes during vaginaldelivery is high in neonates exposed to asymptomatic shedding[220–223], since serial HSV genital cultures during the last fewweeks of gestation are no longer recommended as a methodto prevent neonatal herpes. Instead, women are examined at thetime of labour and caesarean section is carried out only if there isan identifiable lesion. The use of fetal scalp electrodes may alsoincrease the risk for neonatal infection [224,225]. HSV infec-tions should be suspected as the cause of any vesicle appearingin the neonate.

5.1.2. Laboratory assessment of HSV infection inpregnancy and in neonates

Clinical symptoms compatible with genital herpes duringpregnancy require laboratory assessment (Fig. 4). Diagnosis ofHSV infection relies on both serological and virological meth-ods, but the diagnostic process may be complicated due to thenature of the viral infection. Recurrent infections and reacti-vd

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.1. Introduction

.1.1. The pathogenHerpes simplex virus (HSV) establishes latent infection fol-

owing primary infection which may lead to reactivation. It isneurotropic member of the herpesvirus family and the genus

onsists of two types: HSV type 1 (HSV-1) and HSV type 2HSV-2).

The consequences of infection with HSV can vary fromsymptomatic to the life-threatening diseases manifested asSV-encephalitis and neonatal herpes [211]. Recurrent infec-

ions with one type after primary infection with the other typere common. HSV-1 is usually transmitted through the oral routend causes disease in the upper part of the body, while HSV-2 is aexually transmitted virus which tends to cause primarily genitalerpes. However, both HSV-1 and HSV-2 can cause genital her-es which can be a severe disease in primary episodes [211–214].n increase in the prevalence of genital herpes infection has beenocumented worldwide: HSV-2 was the main cause of genitalerpes during the 1980s but since the 1990s HSV-1 constitutesn increasing proportion of the cases. This shift from HSV-2 toSV-1 may have implications for prognosis [215]. Assessmentf risk and diagnosis of HSV infections may involve testing foroth HSV-1 and HSV-2.

HSV infection has serious consequences for the fetus andeonate. Before 20 weeks of gestation, transplacental trans-ission can cause spontaneous abortion in up to 25% of the

ases [216,217]. Later in pregnancy HSV infections are notssociated with increase in spontaneous abortions but intrauter-ne infections may occur. Symptomatic and asymptomatic firstpisodes of genital herpes but not asymptomatic recurrent

ations pose a special challenge to serology, and therefore viraletection in genital lesions is more reliable as a diagnostic mean.

The standard laboratory method to confirm current HSVnfection is virus isolation and typing in cell culture since HSVrows readily in tissue culture. The virus may be isolated within–4 days from swabs taken from herpetic skin and laryngeal orenital lesions (Fig. 4).

PCR techniques improved the correct diagnosis of HSVnfections especially in cases without clear overt manifestations226–229] and can detect viral DNA from lesions that are cultureegative. For genital swabs it might be too sensitive to reflectrue reactivation, and it is not clear whether a positive PCR in

patient with a negative HSV culture reflects a true risk ofransmission of the virus. Therefore physicians should interpretesults with caution and according to additional criteria. Supple-ental serological testing can be used when genital lesions are

ot apparent.In primary infections antibodies appear within 4–7 days after

nfection and reach a peak at 2–4 weeks. Antibodies persistor life with minor fluctuations. Specific IgM antibodies appearfter primary infection but may be detectable during reccurentnfections as well [226]. HSV-1 and HSV-2 share cross reactivepitopes of the surface glycoproteins which are the major targetsf serum antibodies. Therefore it is difficult to identify a newlycquired infection with one HSV type on the background ofre-existing immunity to the other type (usually infection withSV-1 precedes infection with HSV-2). Type specific serologi-

al assays were developed [230] which may be valuable for theanagement of a pregnant woman and her partner: these assays

an identify previous infections, seroconversions and discordantouples. The results may provide information to the pregnant


woman about her risk of acquiring and transmitting HSV to herinfant (Fig. 4). The cost-benefit value of general screening fortype-specific antibodies in pregnant women and their partnerswere recently assessed [231,232].

Rapid and sensitive diagnosis of neonatal HSV diseaseis of utmost importance for initiation of acyclovir treatmentand improvement in the outcome of neonatal herpes has beenachieved due to the application of PCR as a diagnostic tool [233].Neonatal HSV infections are confirmed through positive viralculture or PCR from skin, eye lesion, nasopharynx, rectum orCSF and detection of IgM in a neonate is highly significant[234]. Neonatal HSV infection can occur in newborns withoutthe presence of any vesicular skin disease [229,235,236].

5.2. Laboratory assays for assessment of HSV infection andimmune status

5.2.1. HSV IgG assaysSeveral techniques are used for detection of HSV specific

IgG, among them CF, NT, IF and ELISA. The principles ofthese assays were described above for VZV and are shown inTable 1. There are assays which do not distinguish between IgGdirected to HSV-1 or HSV-2. These assays are useful in primaryinfection in the absence of prior immunity, and they use crudepreparations of cells infected with HSV-1 and/or HSV-2. Theymay detect infection with HSV-2 on the background of HSV-1o

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for HSV is the most accurate method for type-specific serology[241]. This method is the serology gold standard and is per-formed only by few laboratories. WB is expensive to performand requires 2–5 days for completion. New approaches to anti-gen preparation include the application of recombinant gG-1 andgG-2 from a mammalian expression systems to nitrocellulose,or purification of glycoprotein gG2 by lectin chromatography[242].

5.2.3. HSV IgM assaysThese assays are based primarily on ELISA and IF methods

as described for VZV. Due to the persistent nature of the viralinfection and the frequency of recurrent infections, the presenceof IgM is of high value primarily in neonates and infants. In mostcases testing for IgM must be accompanied by testing for IgG,since a negative IgM result does not rule out recent infection.

5.2.4. Virus isolation in tissue cultureHSV is a labile virus, and successful virus culturing from

clinical specimens depends on appropriate sample collection andmaintenance of the cold chain. Swabs should be placed into viraltransport medium and other samples should be placed in a sterilecontainer. Rapid transportation of specimens to the laboratoryand avoiding freeze-thawing cycles results in more than 90%sensitivity of culturing from skin lesions. However, poor samplequality due to inappropriate sampling or compromised transportcaaV1[wp

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[237]. More accurate type-specific assays were developed inrder to detect recurrent infections with either type.

.2.2. HSV type-specific IgG assaysType specific serological assays vary in their abilities [230].

T antibody level can be measured specifically for each type byhe plaque reduction assay using serial dilutions against a fixedose of HSV-1 or HSV-2, with 50% reduction in the plaqueumber as an endpoint. However these assays are accurate onlyn patients infected with only one virus type. NT assays weresed for determination of HSV-2 infection based on the ratioetween the antibody titers to the two viruses. Positive casesere those with HSV-2 to HSV-1 ratio of ≥1 [237,238].IF can be used for detection of antibodies to HSV cell sur-

ace and internal proteins. IF correlates well with NT titers butubtyping is difficult. ELISA assays utilize a variety of condi-ions and antigens. These tests are more sensitive than NT andF and can detect antibodies within the first week of infection.

anufacturers of those tests use mathematical calculations tonfer the presence of HSV-1 or HSV-2 antibodies based on theirelative ratios. New type-specific tests, which distinguish HSV-from HSV-1 are based on glycoproteins gG1 and gG2 [239]

nd can provide useful information on the etiology of the genitalerpes in the symptomatic patient when viral culture and PCRre not helpful. These tests allow the identification of patientsith unrecognized genital herpes [240,241]. There are few good

ommercial ELISA kits for type specific serology which werepproved by the FDA. Nevertheless, HSV type specific antibodyests are interpreted in the context of the clinical history, clin-cal presentation and other laboratory test results for herpes orther possible etiological agents. Finally, Western blotting (WB)

onditions may reduce sensitivity by 20% for primary episodesnd down to 50% for recurrent episodes even if herpetic lesionsre present. When an appropriate sensitive cell line (such asero cells) is used, appearance of a characteristic CPE within8–96 h after inoculation points to the presence of active HSV237]. Direct immuonofluorescence detection of HSV in culturesith CPE is regarded as the standard for confirmation of HSVresence.

The so-called “Shell vial” method, described earlier for CMVsolation (chapter 2.5 and Table 1) shortens the time requiredo identify HSV in specimens to 24–48 h. An innovated com-

ercially available diagnostic test for rapid detection of HSV inissue culture is the ELVIS HSV test kit. It utilizes a recombinantell system in which a reporter enzyme is quickly accumulatednside the infected cells and is detected by intense blue color. Theest is sensitive and specific and detects both HSV-1 and HSV-2243]. The kit is expensive and is used only by few laboratories.

.2.5. Direct antigen detection of HSVOther rapid assays such as the direct fluorescent assay (DFA)

r enzyme immunoassays (EIA) performed directly on smearedells recovered from lesions are rapid but less sensitive thanirus culturing. Accordingly, for detection of virus in genitalwabs they may have similar sensitivity to viral culture forymptomatic shedding but have reduced sensitivity for detectingsymptomatic viral shedding.

.2.6. Detection of HSV DNA by PCRMethods for detection of HSV DNA, particularly PCR tech-

iques (PCR, nested PCR and real-time PCR) were describedbove for other viruses. They are highly sensitive especially for


samples with very low concentration of virus, as was demon-strated by numerous research studies. The performance of PCRanalysis is dependent upon the quality of the specimen. Thereported sensitivity for PCR is 75–100% for the diagnosis ofCNS HSV infection [228,244,245]. The reason for the broadrange in sensitivity is the differences in methodologies andvariability in performance of PCR between laboratories. Stan-dardization using reference samples to assure identical resultsis not yet in place [227].

5.3. Summary

HSV-2 is the main type involved in congenital and neonatalinfection, although genital HSV-1 infections have become morecommon in recent years. Primary maternal symptomatic andasymptomatic infections may cause a serious risk of abortionand fetal growth retardation, while infection during delivery maycause a devastating neonatal disease.

Laboratory assessment of maternal infection and immune sta-tus as well as neonatal infection is critical for timely managementand treatment. Virus isolation from genital lesions is the primarytool for assessment of maternal infection. Serological asays areuseful but are complicated by pre-existing heterotypic antibod-ies. Type-specific IgG assays must be applied for determinationof recurrent maternal infections.

Virus isolation, antigen detection or DNA detection by PCRid

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[261,262] and fetal infection may lead to severe anemia, general-ized edema, congestive heart failure, myocarditis, fetal hydropsand fetal death [263–267]. Fifty percent of women at childbear-ing age are susceptible to parvovirus B19 infection as judgedby the lack of IgG antibodies to the virus [267–272]. How-ever, transplacental transmission rates have been estimated atbetween 25% and 33%, and the incidence of fetal loss is con-sidered between 1.66% and 9% [271,273]. Since intrauterineblood transfusions can increase fetal survival rate if done earlyenough, it is critical to diagnose maternal and fetal infectionaccurately and timely, and laboratory testing is an essential partof the diagnosis [274–279].

6.1.2. Laboratory assessment of parvovirus B19 infectionin pregnancy

Laboratory diagnosis of parvovirus B19 infection has beencomplicated by the nature of the virus, the viral infection pro-cess and the immune response. Parvovirus B19 is a fastidiousvirus which cannot be grown in regular continuous cell lines. Inthose cells where it grows it replicates poorly and the virus yieldis minimal, thus viral culturing is not a diagnostic option andcannot be used for NT assays or for preparation of native viralantigens for serology [261,280,281].

Assessment of parvovirus B19 infection in pregnant womenrelies primarily on serology (Fig. 5). During acute infection IgMantibodies appear 7–14 days from infection with Parvovirus B19asdvor[lcb

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n skin lesions samples are additional tools for rapid and sensitiveiagnosis of symptomatic current infection.

Prenatal diagnosis is not common and has not been assessedor sensitivity and specificity. Neonatal infection is diagnosedy virus culturing or PCR in skin lesions or other clinical speci-ens in case of a disseminated disease, and by detection of IgM

ntibodies.An algorithm describing the laboratory diagnostic assays for

SV infection in pregnant women and neonates is shown inig. 4.

. Parvovirus B19

.1. Introduction

.1.1. The pathogenHuman parvovirus B19 (B19) can infect humans at all ages.

t is a member of the family Parvoviridae, genus Erythrovirushich comprise small, non-enveloped viruses containing linear

ingle-stranded DNA genomes. Most of the humoral immuneesponse is directed to the two capsid proteins VP1 and VP2246]. The seroprevalence is 50% by age 15 and exceeds 80% byge 70 [247]. Childhood infection is mild leading to the diseaserythema infectiosum (“fifth disease”) [248–251] In adults thelinical symptoms vary from asymptomatic or mild to severellness including exanthematous disease, arthropathies, aplasticrisis and, in immunocompromised patients, to chronic anemia252–260].

Maternal infection during pregnancy may lead to fetal infec-ion since viremia is very high and lasts for 6–8 days. Theirus replicates in erythroid precursor cells and fetal tissues

nd may last for 6 month or even longer. IgG antibodies appeareveral days after IgM and persist for life. The antibodies pro-uced specifically react with both structural and non-structuraliral proteins and recognize linear and conformational epitopesf the capsid proteins VP1 and VP2 [282–288]. The antibodiesecognizing conformational epitopes last longer for both IgM285] and IgG [282,284,286], and IgG antibodies recognizinginear epitopes disappear around 6 month post infection. Thisreates technical problems for serological assays as describedelow.

Maternal infection during pregnancy is frequently asymp-omatic and by the time of fetal infection and symptoms, 2–12eeks after maternal infection, she may have high IgG ando IgM responses [260,275,289,290]. Viral DNA detection byighly sensitive molecular assays such as dot-blot hybridiza-ion and PCR can be applied to maternal serum or whole blood276,291]. However, parvoviruses are known for their ability tontegrate into the host genome and to establish persistent infec-ions [292–296]. Recent studies [297–299] showed that B19NA can be detected in serum samples by highly sensitive meth-ds up to 6 months after acute phase viremia, although quanti-ative assays showed that the amount of DNA present decreasedith time [300]. It has also been shown that 0.01–0.03% of theealthy blood donors have B19 DNA detected in their serum301–303]. Thus, detection of B19 DNA in maternal blood byCR might not be related to recent infection.

.1.3. Prenatal laboratory assessment of congenital B19nfection

In many cases congenital B19 infection is suspected onlyhen typical fetal abnormalities are observed. Since the fetus


does not always develop IgM it is necessary to detect the virusitself in fetal samples. B19 cannot be cultivated and thereforedetection of viral DNA is sought. The type of specimen andthe detection methods are still controversial: fetal blood and AFare the most common specimens obtained and the most highlysensitive molecular methods are employed. However, 10–25%of the specimens from asymptomatic fetuses may have B19 DNA[289,290]. This drives the need to develop quantitative molecularassays which can determine viral load and possibly associateit with the risk to the fetus. Those assays include quantitativePCR-ELISA, in situ hybridization and rt-PCR. The latter hasadvantages which will be discussed below [304–308].

6.2. Laboratory assays for assessment of parvovirus B19infection

6.2.1. B19 IgM and IgG assaysSeveral commercial tests have been developed on the basis of

recombinant and synthetic antigens [309–311]. These recombi-nant antigens frequently lack important conformational epitopsreducing the sensitivity and specificity of the assays. More-over, “gold standard” methods like virus NT or HI assays areabsent, and it is difficult to assess the sensitivity and specificityof the serological assays. The variation in specificity and sensi-tivity among commercial assays is high, and discordant resultsare often obtained indicating false positive and false negativer

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serum samples from blood donors with no IgM were tested.Overall, 353 sera were found positive by all methods combined.Of those 98.6%, 94.6% and 89% were positive by assays a, band c, respectively. Some sera reacted only with conformationalepitops. The results of this study underscore the need to useELISA-IgG assays which include conformational epitopes ofboth VP1 and VP2 or VP2 alone, and demonstrate the lowersensitivity of the Western blot assay.

In conclusion, when evaluating recent or past infections inpregnant women it is important to use well-established ELISAassays, and in cases of discrepancy between the test results andthe clinical and epidemiological circumstances, confirmationmust be sought using other commercial assays as well as sup-portive molecular assays discussed next.

6.2.2. Detection of viral DNA in maternal and fetalspecimens

Due to the limitations of the serological assays describedabove and since viremia is long-lasting in B19 infections, molec-ular assays for detection of B19 viral DNA were developed early.The methods most frequently used are DNA hybridization andPCR.

Molecular hybridization assays employing DNA probesderived from most of the viral genome are labeled with 32P,biotin or digoxigenin [319–322] and are capable of detectingapproximately 104–105 genome copies/ml [275,323]. This assayivbPsbni1umtosahpisab

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esponses [314–317].Studies aimed at evaluating the sensitivity and specificity of

arious such assays were mostly based on comparative evalu-tion, using samples selected on the basis of clinical diagno-is. In a study conducted in Norway [315], five commercialLISA and IF IgM kits were comparatively evaluated in threeroups of patients. The calculated specificities for the kits were0.1–94.8%, but the authors refrained from determining sensi-ivity because of the absence of suitable reference methods. Innother study conducted in Sweden [316], four commercial IgMssays were evaluated in comparison to an IgM antibody captureadioimmunoassay as a reference method [318]. The calculatedensitivity varied between 90% and 97% and the specificitiesaried from 88% to 96%.

In a third study conducted in the USA [314] three ELISA sys-ems utilizing one or more conformational antigens for detectionf B19 IgM or IgG in sera of 198 pregnant women were compar-tively evaluated. Agreement with the consensus results variedrom 92.3% to 100% for IgM and from 97.9% to 99.5% for IgG.he authors note the high agreement in this study compared to

heir previous study [312] relating to the fact that the antigenstilized in the two studies were substantially different: the ear-ier study used a linear VP1 antigen while the latter one used aonformational VP2 antigen.

A study designed to evaluate the most appropriate assays forgG detection was conducted in Italy [313] which compared theerformance of three commercial assays using different anti-ens: (a) an ELISA assay using VP1 + VP2 recombinant nativeonformational antigens, (b) an ELISA assay using VP2 recom-inant native conformational epitopes, (c) a Western blot assaysing denatured linear antigens. Four hundred and fourty six

s sensitive enough to detect viral DNA in serum during peakiremia since the viral load exceeds 1010 genome copies/ml,ut low levels of viremia may be missed [275,296,298]. TheCR assays, developed later relied on a variety of primerets derived from different genomic regions and were capa-le of detecting 102–105 genome copies/ml [324,325]. Theested PCR, the PCR-ELISA and the PCR-hybridization assaysncreased sensitivity and moved the detection limit down to–10 genome copies/ml [291,292,296,297,299,326–328]. Bysing those highly sensitive methods it was revealed that inany cases serology failed to detect maternal or fetal infec-

ions, as some seronegative mothers as well as fetuses turnedut positive in the DNA detection assays. This finding was noturprising in view of the known limitations of the serologicalssays. The ability to detect maternal and fetal infection by theighly sensitive molecular methods allowed correct diagnosis ofreviousely unresolved cases, particularly cases of intrauterinenfection which did not result in fetal hydrops or were completelyubclinical [275,276,290,292,296,297,324,327,329]. However,s stated above, low levels of B19 DNA in maternal blood cane unrelated to recent infection.

The use of various techniques without standardization, thepidemiologically variable circumstances (epidemic VS non-pidemic years), and the use of different primer sets and clinicalpecimens led to a high variability in the sensitivity, specificitynd interpretation of the assays results. Moreover, new geno-ypes discovered recently were missed by common primer sets.omparison of IgM and DNA detection by PCR in fetal bloodas conducted in a study done on 57 pregnant women and their

etuses who had abnormal ultrasonography [327]. Viral DNAas found by PCR in 16 out of 58 fetuses (27%) while IgM was


detected only in 7 (12.3%). Two fetuses had false-positive IgMresult, not supported by any other findings. Other researchers[276] compared maternal IgM to fetal serum or AF PCR resultsin 56 women at high risk for B19 infection. They found pos-itive PCR in 24 IgM-negative/IgG-positive and 4 seronegative(total 50%) out of, in addition to positive PCR in 15 (26%) ofIgM-positive women. Another group [282] reported detection ofviral DNA in fetal serum or AF by a PCR-hybridization assayin 11 out of 80 cases of fetal hydrops (14%) while maternal IgMantibodies were detected only in 3 (3.7%). Finally, a compre-hensice prospective study in 18 fetal hydrops cases conductedin Italy [275] examined maternal serum, fetal cord blood andamniotic fluid using nested PCR, dot-blot hybridization and insitu hybridization (ISH). The results showed that the ISH assayin fetal blood cells was 100% sensitive while the other methodsmissed few to many cases. The conclusion drawn from this studyis that various assays have complementary roles, and reliablediagnosis can be achieved only by a combination of serologi-cal and molecular assays done on maternal and fetal samples asoutlined in Fig. 5.

6.2.3. Quantitative assays for detection of viral DNAThe need to determine the quantity of B19 viral DNA present

in clinical samples (viral load) arose because B19 can establishlong-lasting persistent infection in immunocompetent individ-uals. It is not clear if low viral loads reflect whole genomesoSriietov

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6.3. Summary

Parvovirus B19 infection during pregnancy may cause fetaldamage resulting in fetal loss. Early diagnosis of maternal infec-tion will allow fetal assessment and treatment by intrauterineblood transfusion. Unfortunately, mothers often are not awareof their infection until fetal damage is observed.

Confirmation of B19 infection requires laboratory assess-ment, which is complicated by the nature of the viral infectionand immune response. Serology, performed by using ELISAassays rely on recombinant antigens and concordance is lowamong all commercial assays available. In the absence of a “goldstandard” assay false positive and false negative results prevail.Virus culturing is impossible and virus detection is based onvarious molecular assays.

In spite of several studies there is no consensus regarding themost appropriate clinical specimen and method for detection ofviral DNA. Currently, on practical grounds, it is recommendedto use ELISA IgM and IgG assays based on recombinant con-formational epitopes of VP1 and VP2 or VP2 alone, and to useAF or fetal serum for detection of fetal infection by the mostsensitive molecular methds available (nested PCR or rt-PCR).Since B19 may establish long lasting infection in the absenceof symptoms, interpretation of viral DNA detection in mater-nal blood is difficult. Assessment of fetal infection and riskshould rely on the clinical situation and other prenatal diagnosticm

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r viable infectious virus particles, or only pieces of viral DNA.tudies in seronegative plasma-pool recipients showed that onlyecipients of plasma containing >107 genome copies/ml becamenfected or seroconverted [305]. B19 DNA can also be detectedn solid organ tissues by PCR for years [323,330,331]. As notedarlier viral DNA can be detected in fetal tissue or blood inhe absence of any detectable congenital abnormalities, and theutcome of detectable fetal infection, including fetal hydrops,aries from spontaneous resolution to still-birth.

Real-time PCR assays developed recently are equivalent tor more sensitive than nested PCR and PCR-ELISA but areuch less prone to molecular contamination and produce a

uantitative result which can be standardized and automated304,306–308,332]. In a retrospective study, Knoll et al. [332]sed real-time PCR to investigate the viral load in paired samplesrom mothers and their abnormal fetuses, and from mothers withormal fetuses who were exposed to B19. The viral load in theaternal serum ranged from 7.2 × 102 to 2.6 × 103. The authors

id not report statistically significant correlation between mater-al or fetal viral load and fetal condition. However, they notedhat they could not exclude a correlation between peak mater-al viremia levels and fetal condition since most of the mothersere not aware of their infection until onset of fetal symptoms,

nd their serum was collected after peak viremia. Although theiral load in fetal sera was higher than in AF samples, the rt-CR assay detected all positive cases using either one of theseetal specimens. The authors recommend testing AF rather thanetal blood because AF can be drawn earlier, is simpler to obtainnd less risky to the fetus. Clearly, more prospective studies areecessary on larger groups of patients to learn more about thessociation between viral load and pregnancy outcome.

eans.An algorithm describing the most practical approach to lab-

ratory assessment of B19 infection in pregnancy is shown inig. 5.

. Human immunodeficiency virus

.1. Introduction

.1.1. The pathogenHuman immunodeficiency virus (HIV) is a retrovirus that

nfects helper T cells of the immune system causing a progressiveeduction in their numbers, and eventually acquired immunode-ciency syndrome (AIDS). HIV is a member of the Retroviridaeamily [333], genus Lentivirus (or “slow” viruses). The coursef infection with these viruses is characterized by a long intervaletween initial infection and the onset of serious symptoms. Theingle-stranded RNA viruses exploit their reverse transcriptasenzyme to synthesize DNA using their RNA as a template. TheNA is then incorporated into the genome of infected cells.AIDS was first diagnosed in European sailors with African

onnections [334–336]. The first AIDS virus, HIV-1, was ini-ially identified in 1983 [337–341]. A second AIDS virus, HIV-2,as discovered in 1986 [342–344]. Forty million people were

stimated to be infected with HIV at the end of 2004 [345]. Theighest prevalence is found in Sub-Saharan Africa and it is risingainly in Asia and some of the former Soviet Union countries

ike Ukraine and the Russian Federation [345,346]. During itspread among humans, group M HIV-1 (one of three groups: M,

and M), has evolved into multiple subtypes that differ fromne another by 10–30% along their genomes [347–350].


HIV is passed on primarily via four routes: unprotectedsexual intercourse (both homosexual and heterosexual), shar-ing of needles by IV drug users, medical procedures usingHIV-contaminated blood, tissues or equipment, and mother-to-child transmission (MTCT). The likelihood of transmissionis increased by factors that may damage mucosal linings ofexposed tissues, especially by other sexually transmitted dis-eases that cause ulcers or inflammation.

HIV can be transmitted from infected mothers to infants dur-ing pregnancy (intrauterine) through transplacental passage ofthe virus [351], during labor and delivery (intrapartum) throughexposure to infected maternal fluids (blood or vaginal secretions)[352–355] and during the post partum period through breast-feeding [356–359]. Infants who have a positive virologic test(see below) at or before age 48 h are considered to have early(i.e., intrauterine) infection, whereas infants who have a negativevirologic test during the first week of life and subsequent posi-tive tests are considered to have late (i.e., intrapartum) infection[360]. In the absence of breast-feeding, intrauterine transmissionaccounts for 10–35% of infection, and 60–75% of transmissionoccurs during labor and delivery [361–363]. Without interven-tion, maternal infection leads to about 25–30% of babies beinginfected [364–369].

Use of anti-retroviral therapy (ART) during pregnancy inHIV-infected women is associated with improved obstetric out-come of reduced infection rates of babies to less than 2%[Uhpaastr

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Knowledge of maternal HIV infection during the antena-tal period enables HIV-infected women to receive appropriateantiretroviral therapy during pregnancy, during labor, and tonewborns to reduce the risk for HIV transmission from motherto child [370,383,384]. It also allows counseling of infectedwomen about the risks for HIV transmission through breastmilk and advising against breast feeding in countries where safealternatives to breast milk are available [385]. Early diagnos-tic evaluation of HIV-exposed infants permits early initiationof aggressive antiretroviral therapy in infected infants and ini-tiation of prophylaxis against Pneumocystis carinii pneumonia(PCP) in all HIV-exposed infants beginning at age 4–6 weeks inaccordance with Public Health Services (PHS) guidelines [386].

7.1.3. Prenatal laboratory assessment of HIV infectionPrenatal laboratory assessment of congenital HIV infection in

AF or cord blood is not recommended as the invasive proceduresincreases the risk of transmitting the virus from the maternal tothe fetal blood stream.

7.2. Laboratory assessment of HIV infection

7.2.1. HIV antibody assaysThe initial screening for HIV infection in adults and chil-

dren is done by testing for antibodies. Viral load assays arenot intended for routine diagnosis but could be used in clini-ccdithddittw

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370–373] and little maternal toxicity [374–376]. The currentS guidelines are to offer all pregnant HIV-1-infected womenighly active antiretroviral therapy (HAART) to maximally sup-ress viral replication, reduce the risk of prenatal transmission,nd minimize the risk of development of resistant virus. Inddition, HIV-infected women are offered an elective caesareanection delivery [355]. The results also suggest that special atten-ion should be given to women belonging to previously identifiedisk groups.

Because testing has proven very successful in helping to pre-ent the spread of the disease to babies, a US federal panel hasecommended that all pregnant women, not just those consid-red at high risk, be screened for the AIDS virus [377–381]. Norenatal (intrauterine) diagnosis is performed because: (a) thenfection can occur during delivery and (b) the testing interven-ion may increase the chance of virus transmission to the baby.ssessing the infection status of the mother is critical [381] and

ollowing delivery the new-born should be tested. The laboratoryesting is essential for the diagnosis.

.1.2. Importance of laboratory assessment of HIVnfection in pregnancy

Identification of HIV-infected women before or during preg-ancy is critical to providing optimal therapy for both infectedomen and their children and to preventing perinatal trans-ission (see below). For women with unknown HIV status

uring active labor, ART can still be effective when given duringabor and delivery, followed by treatment of the newborn [382].his expedited intervention requires the use of rapid diagnos-

ic testing during labor or rapid return of results from standardesting.

al management of HIV-infected persons in conjunction withlinical signs and symptoms and other laboratory markers ofisease progression. Detection of HIV-1 p24 antigen (Table 1)s used for routine screening in blood and plasma centers butheir routine use for diagnosing HIV infection in individualsas been discouraged because the estimated average time frometection of p24 antigen to detection of HIV antibody by stan-ard enzyme-immuno-assay (EIA) is 6 days, and not all recentlynfected persons have detectable levels of p24 antigen [387]. Inhe USA several FDA approved tests are available that enablehe testing of HIV antibodies in different body fluids such ashole blood, serum, plasma, oral fluid and urine.The standard testing algorithm for HIV-1 consists of initial

creening with an EIA to detect antibodies to the virus. Reactivepecimens undergo confirmatory testing with a more specificupplemental test, usually a Western blot assay (WB) or, lessommonly, IFA (Table 1) [388]. Using both tests increases accu-acy of the results while maintaining their sensitivity [389–391].nly specimens that are repeatedly reactive by EIA and posi-

ive by IFA or reactive by WB are considered HIV-positive andndicative of HIV infection [389,390,392,393].

Incomplete antibody responses that produce negative or inde-erminate results on WB tests can occur among persons recentlynfected with HIV who have low levels of detectable antibodiesi.e., seroconverting), persons who have end-stage HIV dis-ase, and perinatally exposed but uninfected infants who areeroreverting (i.e., losing maternal antibody). Non-specific reac-ions producing indeterminate results in uninfected persons haveccurred more frequently among pregnant women than amongther persons [389–391,394].

False-positive WB results are rare [395].


7.2.2. Detection of viral DNA in maternal and newbornspecimens7.2.2.1. Diagnosis of HIV infection in maternal specimens.Pregnant women are screened for the presence of antibodiesusing the standard testing algorithm for adults (EIA plus WB orIFA) as shown in Fig. 6. For women with unknown HIV statusduring active labor, rapid diagnostic tests are used if rapid returnof results from standard testing is not available.

7.2.2.2. Diagnosis of HIV infection in newborns. Infant HIVtesting should be done as soon after birth as possible soappropriate treatment interventions can be implemented quickly[377,384]. The standard antibody assays used for older childrenand adults are not useful for diagnosing children younger than18 months as the presence of maternal antibodies makes sero-logic tests uninformative. Therefore, a definitive diagnosis ofHIV infection in early infancy requires viral diagnostic assays,including HIV-1 p24 antigen assays, nucleic acid amplification(e.g., PCR) or viral culture. HIV infection can be definitivelydiagnosed in most infected infants by age 1 month and in vir-tually all infected infants by age 6 months. HIV infection isdiagnosed by at least two positive assays using two separatespecimens [378].

HIV DNA PCR is the preferred virologic method for diag-nosing HIV infection during infancy. It has 99% specificity toidentify HIV proviral DNA in peripheral blood mononuclearcE

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antigen testing alone is not recommended because of its sub-stantiallty reduced sensitivity and specificity compromising thecritical need for timely diagnosis [405].

Whether the current, more intensive antiretroviral combina-tion regimens women may receive during pregnancy for treat-ment of their own HIV infection will affect diagnostic test sen-sitivity in their infants is unknown. Similarly, if more complexregimens are administered to HIV-exposed infants for perinatalprophylaxis, the sensitivity of diagnostic assays will need to bere-examined [401].

7.2.2.3. Different subtypes. HIV subtype B is the predominantviral subtype found in the U.S. and western Europe. Non-subtypeB viruses predominate in other parts of the world, such as sub-type C in regions of Africa and India and subtype E in much ofsoutheast Asia.

Currently the available HIV DNA PCR commercial tests areless sensitive for detection for non-subtype B HIV, and falsenegative HIV DNA PCR assays have been reported in infantsinfected with non-subtype B HIV [406–410]. Caution should beexercised in the interpretation of negative HIV DNA PCR testresults in infants born to mothers who may have acquired an HIVnon-B subtype. Some of the currently available HIV RNA assayshave improved sensitivity for detection of non-subtype B HIVinfection [411–413], although even these assays may not detectsome non-B subtypes, particularly group O HIV strains [414]. IncsonHRPcec1

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ells (PBMC) obtained from whole blood samples collected inDTA-containing tubes [396].

A meta-analysis of published data from 271 infected childrenndicated that HIV DNA PCR was sensitive for the diagnosis ofIV infection during the neonatal period. Thirty-eight percentf infected children had positive HIV DNA PCR tests by age8 h, 93% by age 14 days and 96% by age 28 days. No sub-tantial change in sensitivity during the first week of life wasbserved, but sensitivity increased rapidly during the secondeek.Quantitative assays that detect HIV RNA in plasma (see Sec-

ion 7.2.2.3 below) appear to be as sensitive as HIV DNA PCRor early diagnosis of HIV infection in HIV-exposed infants397–402]. The specificity is comparable between the two tests,ut results of HIV RNA load below 104 copies/ml should benterpreted with caution [403]. Some clinicians use HIV RNAssay as the confirmatory test for infants testing HIV DNA PCRositive since it provides viral load measurement which guidesreatment decisions. Available quantitative RNA tests includehe Amplicor HIV-1 monitor test 1.5 (Roche Diagnostics), theASBA EasyQ HIV-1 (BioMerieux), the Quantiplex HIV RNA.0 (bDNA) (Bayer) and the LCx HIV RNA quantitative assayAbbott Laboratories) assays [416–421]. However, special atten-ion should be taken where non-B HIV is expected.

HIV culture for the diagnosis of infection has a sensitivityhat is similar to that of HIV DNA PCR [404]. However, HIVulture is more complex and expensive to perform than DNACR, and definitive results may not be available for 2–4 weeks.

Both standard and immune-complex-dissociated p24 anti-en tests are highly specific for HIV infection and have beensed to diagnose infection in children. However, the use of p24

ases of infants where non-subtype B perinatal exposure may beuspected and HIV DNA PCR is negative, repeat testing usingne of the newer RNA assays shown to be more sensitive foron-subtype B HIV is recommended (for example, the AmplicorIV-1 monitor test 1.5, Nuclisens HIV-1 qt or Quantiplex HIVNA 3.0 (bDNA) assays). In children with negative HIV DNACR and RNA assays but in whom non-subtype B infectionontinues to be suspected, the clinician should consult with anxpert in pediatric HIV infection and the child should undergolose clinical monitoring and definitive HIV serologic testing at8 months of age.

.2.2.4. Test algorithm for neonates. HIV infection is diag-osed by two virological tests performed on separate bloodamples, regardless of age. The testing rules are summarizedelow and shown in Fig. 6:

1) Initial testing is recommended by age 48 h. As many as40% of infected infants can be identified at this time. Bloodsamples from the umbilical cord should not be used for diag-nostic evaluations, because of concerns regarding potentialcontamination with maternal blood.

2) Repeated diagnostic testing can also be considered at age14 days in infants with negative tests at birth. The diag-nostic sensitivity of virological assays increases rapidly byage 2 weeks. Early identification of infection would permitdiscontinuation of neonatal ZDV chemoprophylaxis and afurther evaluation of the need for more aggressive drug com-bination therapy.

3) Retest infants with initially negative virological tests at age1–2 months. Using ZDV monotherapy to reduce perinatal


transmission did not delay the detection of HIV in infantsin PACTG protocol 076 [370,400–402,415].

(4) At age 3–6 months retest HIV-exposed children who havehad repeatedly negative virological assays at birth and atage 1–2 months.

(5) HIV infection can be reasonably excluded in non-breast fedinfants with two or more negative virologic tests performedat age >1 month, with one of those being performed at age>4 months [386].

(6) Two or more negative HIV immunoglobulin G (IgG) anti-body tests performed at age >6 months with an interval of atleast 1 month between the tests can also be used to reason-ably exclude HIV infection in HIV-exposed children with noclinical or virologic laboratory evidence of HIV infection.

(7) Serology after 12 months is recommended to confirm thatmaternal HIV antibodies transferred to the infant in uterohave disappeared if there has not been previous confirmationof two negative antibody tests.

(8) If the child is still antibody-positive at 12 months, then test-ing should be repeated between 15 and 18 months [387].Loss of HIV antibody in a child with previously negativeHIV DNA PCR tests definitively confirms that the child isHIV uninfected.

(9) A positive HIV antibody test at >18 months of age indicatesHIV infection [378].

7

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[2] Melnick JL. Taxonomy of viruses. Prog Med Virol 1976;22:211–21.

[3] Oker-Blom C, Kalkkinen N, Kaariainen L, Pettersson RF. Rubella viruscontains one capsid protein and three envelope glycoproteins, E1, E2a,and E2b. J Virol 1983;46:964–73.

[4] Vesikari T. Immune response in rubella infection. Scand J Infect Dis1972;4:1–42.

[5] Mitchell LA, Zhang T, Ho M, Decarie D, Tingle AJ, Zrein M, et al.Characterization of rubella virus-specific antibody responses by usinga new synthetic peptide-based enzyme-linked immunosorbent assay. JClin Microbiol 1992;30:1841–7.

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[8] Coulter C, Wood R, Robson J. Rubella infection in pregnancy. Com-mun Dis Intell 1999;23:93–6.

[9] Webster WS. Teratogen update: congenital rubella. Teratology1998;58:13–23.

[10] Horstmann DM, Schluederberg A, Emmons JE, Evans BK, Ran-dolph MF, Andiman WA. Persistence of vaccine-induced immuneresponses to rubella: comparison with natural infection. Rev InfectDis 1985;7(Suppl. 1):S80–5.

[11] Harcourt GC, Best JM, Banatvala JE. Rubella-specific serum andnasopharyngeal antibodies in volunteers with naturally acquired andvaccine-induced immunity after intranasal challenge. J Infect Dis1980;142:145–55.

[12] Just M, Just V, Berger R, Burkhardt F, Schilt U. Duration of immunityafter rubella vaccination: a long-term study in Switzerland. Rev InfectDis 1985;7(Suppl. 1):S91–4.

[13] Balfour Jr HH, Amren DP. Rubella, measles and mumps antibodies
.3. Summary
Identification of HIV-infected women before or during preg-ancy is critical to providing optimal therapy for both infectedomen and their children and to preventing perinatal transmis-

ion. Pregnant women are screened for the presence of antibod-es using the standard testing algorithm for adults (EIA plus WBr IFA). Prenatal laboratory assessment of congenital HIV infec-ion is not recommended as it increases the risk of infecting theetus, but extensive testing is performed to assess the infectiontatus of the baby following delivery. A definitive diagnosis ofIV infection in early infancy requires repeated testing usingirological assays, with HIV DNA PCR being the currently pre-erred method. All infected infants can be definitively diagnosedy age 6 months.

An algorithm describing the laboratory diagnostic assays forIV infection in pregnant women and neonates is shown inig. 6.

cknowledgments

We are grateful to Galit Zemel for her extensive technical sup-ort in organizing the citations and reference list and in preparinghe review for submission. We also thank Zehava Yosefi for help-ng in reference retrieval and typing.

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Laboratory assessment and diagnosis of congenital viral infections